EXERCISE 1 - BrainMass
LABORATORY EXERCISES IN GENETICS
Melvin Beck
University of Memphis
May 2004
TABLE OF CONTENTS
Safety Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .3
Exercise 1: Examination of Wild-type and Mutant
Drosophila melanogaster . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .4
Exercise 2: Competition in a Population Cage . . . . . . . . . . . . . . . . . . . . . . . . . . . 10
Exercise 3: Laboratory Techniques . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14
Exercise 4: Linkage, Crossing Over, and Genetic Mapping in Drosophila . . . . . . .28
Exercise 5: Tetrad Analysis in Sordaria fimicola . . . . . . . . . . . . . . . . . . . . . . . . .33
Exercise 6: Biochemical Genetics: Chromatographic Analysis of
Drosophila Eye Pigments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .42
Exercise 7: Protein Electrophoresis and Hemoglobin Polymorphism. . . . . . . . . . .48
Exercise 8: Restriction Endonuclease Digestion and Analysis of
Lambda DNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .59
Exercise 9: DNA Fingerprinting. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .72
Exercise 10: Transformation: Introduction of Plasmid DNA into
Escherichia coli . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .83
Exercise 11: Bioinformatics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 91
Exercise 12: DNA Amplification by Polymerase Chain
Reaction (PCR) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .. 98
Exercise 13: Lac Operon: Synthesis of ß-Galactosidase in
Escherichia coli . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 107
Appendix A: Instructions for Journal Style Laboratory Reports . . . . . . . . . . . . 114
Appendix B: Instructions for Maintaining a Laboratory Notebook . . . . . . . . . . .120
Appendix C: Regression Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .124
SAFETY PROCEDURES
1. Assume all chemicals are hazardous and avoid any direct contact with them.
2. Always wear appropriate protective equipment. For example, disposable gloves and eye protection should be worn when conducting some of the experiments.
3. Avoid eating, drinking, pencil-chewing, etc. This will minimize the chance that you will ingest hazardous chemicals or materials used in the laboratory.
4. NEVER MOUTH PIPET. Many of the laboratory experiments will involve some type of liquid transfer. These transfers should be done with the appropriate transfer device. It is imperative that you become proficient in the use of these transfer devices.
5. Electrophoresis involves the use of high electrical voltages. Make sure you following the instructions for the proper use of electrophoretic equipment. The power supply should always be turned off before opening the gel apparatus.
6. A ultraviolet transilluminator will be used for visualizing DNA in agarose gels. Make sure you wear UV-blocking goggles, gloves, and a lab coat or long sleeved shirt to protect yourself from the intense ultraviolet light emitted by the transilluminator. This will prevent a severe "sunburn" that can result from even short exposures to the UV light from the transilluminator.
7. Make sure the microcentrifuge is balanced before you spin your samples. Each tube in the rotor of the microcentrifuge should have a corresponding tube on the opposite side of the rotor.
8. Wipe down you work area before leaving the laboratory and always wash your hands before leaving the laboratory.
9. Treat any microbial culture as a potential pathogen. Discard any disposable material that contains bacteria or fungi in a biohazard waste container. Place test tubes and other reusable supplies that contain bacteria in an appropriate container for later autoclaving.
10. Smoking is not permitted in either the genetics laboratory or the Ellington Biology Building.
EXERCISE 1
EXAMINATION OF WILD-TYPE AND MUTANT DROSOPHILA MELANOGASTER
EQUIPMENT
Stereo dissecting microscope
SUPPLIES AND MATERIALS
Various Drosophila melanogaster mutants
Fly-Nap® (Carolina Biological Supply Company)
Camel's hair brushes
Dissecting needles
3 x 5 index cards
Morgue
INTRODUCTION
Transmission of genetic information from one generation to the next is very similar in sexually reproducing, multicellular organisms. Drosophila melanogaster (fruit fly or vinegar fly) is one of the most frequently used organisms in introductory genetics laboratories. D. melanogaster is small, easily handled, and breeds prolifically in the laboratory. The fruit fly has a short life cycle and many mutants are known for this organism.
The fruit fly undergoes complete metamorphosis. The stages of its life cycle are: egg, larva, pupa, and adult (Figure 1). At 20oC the life cycle is completed in about 15 days whereas at 25oC the life cycle takes only 10 days. Lower temperatures prolong the life cycle and impair viability. Higher temperatures increase sterility and reduce viability.
A D. melanogaster females can mate about 6 hours after emerging from the pupa case (puparium). Following mating, females start ovipositing eggs during the second day of adult life. The eggs are white and are laid on the surface of the food. A single female can produce approximately 500 eggs within 2 weeks. The eggs hatch within one day and the larvae feed constantly and burrow through the food, leaving many channels. When the larvae are preparing to pupate, they crawl out of the food and attach themselves to the sides of the culture vial.
Figure 1. Life cycle of Drosophila melanogaster
OBJECTIVE
To learn how to anesthetize, handle, and identify male and female fruit flies. To examine wild-type flies and compare them to various mutants.
ANESTHETIZING TECHNIQUE
1. Never anesthetize stock bottles of flies.
2. Get a clean anesthetizing bottle, stopper, Fly-Nap®, anesthetizing wand, index card, brush, probe, and dissecting scope.
3. Obtain a stock bottle of the desired flies.
4. Gently tap the bottom of the stock bottle with the palm of the hand so flies will fall to the bottom of the vial.
5. Take the anesthetizing bottle and remove its stopper. Hold it upside down over the stock bottle. Take the stopper out of the stock bottle and quickly place the anesthetizing bottle over its lip.
6. Carefully hold the two bottles together with the empty, anesthetizing bottle on top.
7. Several flies should fly into the anesthetizing bottle.
8. Remove the stock bottle. Quickly stopper the anesthetizing bottle and then the stock bottle.
9. Insert an anesthetizing wand into the Fly-Nap® to moisten it and allow any excess liquid on the wand to drip back into the bottle. Push the stopper of the anesthetizing bottle to one side and insert the anesthetizing wand along the side of the stopper so that the anesthetic end is just below the stopper. Keep the vial upright. Do not remove the stopper to insert the anesthetizing wand or until the flies are anesthetized!
10. The anesthetized flies should fall to the bottom of the bottle. Watch the flies closely and remove the stopper and wand when the flies are anesthetized (~1 to 2 minutes). Do not over-anesthetize the flies. Over-anesthetized flies have their wings extended at right angles to the body and are dead.
11. Dump the flies onto an index card and examine them under a dissecting scope. The flies may be picked up using a camel's hair brush. A dissecting needle is useful for segregating flies when counting or sexing them. Anesthetized flies often quiver. If the flies become active before the examination is complete, replace them in the anesthetizing bottle. Over exposure to Fly-Nap® can kill the flies or damage their reproductive ability.
12. Place all discarded flies in the morgue (jar containing water and a household detergent).
SEXING FLIES
It is important to be able to distinguish males from females because males must be chosen from one stock to mate with females of another in the various genetics experiments. The sex of D. melanogaster can be determined in several ways. The following differences are generally used:
1. Sex comb. The sex comb is found only in males and consists of a row of about ten stout, black bristles on the distal surface of the basal tarsal joint of the first pair of legs (Figure 2A).
2. Abdomen. The female has seven visible segments, elongates to a point posteriorly, and has separate dark bands along the dorsal surface to the very tip. The male has five visible segments and has a rounded end posteriorly; dark bands of the last few segments are fused (Figure 2A).
3. Genital region. The female has anal plates and light-colored ovipositor plates. The male has anal plates and darkly pigmented genital arch and penis. Examination of the external genitalia under magnification is the best means of distinguishing the sex of flies (Figure 2B).
4. Size of adult. The female is generally larger than the male (Figure 2A); however, this is not an extremely reliable difference.
COLLECTING VIRGIN FEMALES
Virgin females are needed for controlled crosses because females, which have already mated, will lay eggs fertilized by males of the same stock. Once females have mated they store sperm in a spermathecium and can utilize sperm from a single insemination for most of their reproductive lives. Females are initially sexually immature and do not mate until approximately 6 hours after emerging from the pupae cases. To isolated virgin females, remove all adult flies from the culture bottle, and select females that emerge within the following 6 hours. Females tend to emerge sooner than males raised in the same vial; therefore, more virgin females can be obtained from a vial that has just begun producing adults than from an older culture. In addition, most of the flies will emerge during the early part of the day.
[pic]
Figure 2. Drosophila melanogaster. A. Dorsal view of a female and a male Drosophila. B. Ventral view of the abdomen of a female and a male Drosophila.
CROSSING FRUIT FLIES
The general procedure to be following in establishing a fruit fly cross is:
1. Collect virgin females and cross them with males of a different stock. The cross should be started with at least 5 pairs of flies. The parent flies should be removed 7 days after mating to avoid parents breeding with the F1 flies.
2. Upon emergence of the F1, examine several of them and record their phenotype. Set up two crosses in fresh culture bottles using 5 F1 females and 5 F1 males in each cross. The parents should be removed 7 days after mating.
3. Classify and count the F2 flies as soon after emergence as possible, and at frequent intervals for a period of 10 days after the first flies begin to emerge.
SYMBOLS USED IN DROSOPHILA GENETICS
1. The normal fly is called the wild type, and the genes that yield the normal phenotype are designated by a +. The + may occasionally refer to the entire genome or may simply describe a single gene depending on how it is used in a particular context.
2. The symbol for each mutant allele, however, is derived from the descriptive name of the mutant trait (e.g. vg is used for the vestigial phenotype; Pm is used for the Plum phenotype).
3. Small letters (vg) designates recessive alleles and dominant alleles are indicated by capitalizing the first letter of the symbol (Pm). The wild type gene at the vestigial locus is symbolized as vg+ and is dominant to vg while the wild type gene at the Plum locus is symbolized as Pm+ and is recessive to Pm.
PROCEDURE
1. Examine wild-type flies. Learn how to distinguish males from females.
2. After you are familiar with wild-type flies, examine the various mutant stocks and determine how each mutant differs from the wild-type flies.
SUGGESTED READING
Bennet-Clark, H. and A. Ewing. 1970. The love song of the fruit fly. Sci. Amer. 223:84-92.
EXERCISE 2
COMPETITION IN A POPULATION CAGE
EQUIPMENT
Stereo Dissecting Microscopes
SUPPLIES AND MATERIALS
Population cage
Fly-Nap® (Carolina Biological Supply Company)
Anesthetizing Bottles
Camel's Hair Brushes
Dissecting Needles
Instant Medium Formula 4-24® (Carolina Biological Supply Company)
0.5% Propionic Acid
Wing mutant of Drosophila melanogaster
Wild type Drosophila melanogaster
INTRODUCTION
Population genetics is the study of the genetic composition of populations and how this genetic composition changes from generation to generation. A population is a group of individuals of the same species that are capable of interbreeding and producing viable, fertile offspring. Hardy and Weinberg demonstrated that both gene frequencies and genotype frequencies will remain constant from generation to generation in a large, interbreeding population in which mating is at random and there is no selection, migration, or mutation. Hardy and Weinberg independently devised the formula
p + q = 1
that represents the total of the gene frequency when only two alleles of a gene exist. In the formula, p represents the frequency of one allele and q equals the frequency of the other allele. When this formula is squared and expressed as the binomial (p + q)2 = p2 + 2pq + q2, it can be used to determine the proportion of each genotype in a population at equilibrium. In the binomial, p2 is the proportion of one homozygote, 2pq is the proportion of heterozygotes, and q2 is the proportion of the other homozygote. Thus, the Hardy-Weinberg formula is useful for estimation and comparison of gene and genotype frequencies in a population.
Populations of organisms are not infinitely large, mating may not occur at random, and selection, migration, and mutation do occur. Natural selection is one of the principal factors that alter gene frequencies in populations. Selection refers to the differential reproduction by genotypes within a population. Certain organisms in a population are more fit and will therefore contribute more progeny to the next generation. Thus, the genes they carry are represented at a higher frequency in the gene pool of the next generation.
The fruit fly Drosophila melanogaster is a convenient organism for demonstrating the action of natural selection in populations. Fruit flies of different genotypes can be introduced into a population cage. The persistence or elimination of an allele in the population cage depends on the advantages or disadvantages the allele confers on its carriers in competition with other genotypes. Estimation of gene frequencies in the population cage can be made utilizing the Hardy-Weinberg formula.
OBJECTIVE
To determine the effect of selection on a recessive allele in a population of D. melanogaster.
PROCEDURE
1. Prepare four food cups using instant medium. To prepare the food cups, use 1 small cup of instant medium, 1 small cup of 0.5% propionic acid, and add a pinch of yeast to the top of the instant medium. Do not us an excessive amount of yeast because if the yeast forms a solid layer over the food flies will not lay their eggs. Propionic acid is a mold inhibitor. The food should appear creamy, like mashed potatoes. Food that is too dry will have a flaky appearance. Date the food cups and insert them into the population cage.
2. Place 25 wing mutant females, 25 wild type females, 25 wing mutant males, and 25 wild type males in the population cage. This procedure results in an initial gene frequency of 50% for both the recessive wing mutant gene and the dominant wild type allele. Make sure not to over anesthetize your flies. The wings of anesthetized living flies are folded normally over the abdomen. Flies with wings extended outward from the thorax have been over anesthetized and are dead!
3. Tape the cover securely to the top of the population cage.
4. Label the population cage with the lab section, date, and phenotype of flies.
5. Sample the population cage every 3 weeks. Take a fresh food cup, date the cup, and insert it in place of the oldest food cup. At the next lab meeting, remove the sampling cup and replace it with a freshly prepared food cup.
6. Stopper the sampling cup and label it with your name, lab section, date, and phenotype of the flies.
7. Score and count the adult flies that are produced in the sampling cup for a period of 10 days after the first flies begin to emerge.
8. At the end of the experiment, remove all the food cups and stopper the holes in the population cage. Anesthetize the flies and then score and count the adult flies that are in the population cage.
CALCULATIONS
ALLELE FREQUENCY
The allele frequency (q) for the recessive allele is found by taking the square root of the observed frequency of the recessive homozygote.
q = √q2
From this, the frequency of the dominant allele may be found.
p = 1 - q
GENOTYPE FREQUENCIES
Genotype frequencies may be calculated by substituting the numerical values for p and q in the Hardy-Weinberg formula:
p2 + 2pq + q2 = 1
COEFFICIENT OF SELECTION
The coefficient of selection (s) is the proportion of the recessive homozygotes (q2) eliminated by natural selection from the population in a given generation. Thus, the frequency of the recessive gene in the next generation is
q1 = qo - sq2/1 - sq2
From this equation, we can derive the equation
s = (qo - q1) / qo2(1 - q1)
To find the experimental value for s, substitute the values for qo and q1 obtained in your experiment for any two successive generation in this equation.
SUGGESTED READINGS
Cavalli-Sforza, L. L. 1974. The genetics of human populations. Sci. Amer. 221:30-37.
Crow, J. F. 1988. Eighty years ago: The beginnings of population genetics. Genetics 119:473-476.
Mayr, E. 1977. Darwin and natural selection. Amer. Sci. 65:321-327.
Merrell, D. J. and J. C. Underhill. 1956. Competition between mutants in experimental populations of Drosophila melanogaster. Genetics 41:461-485.
EXERCISE 3
LABORATORY TECHNIQUES
EQUIPMENT
37oC Incubator
Microcentrifuge
Electronic balances
SUPPLIES AND MATERIALS
Pipetting Exercise
1 ml Glass pipettes
Pipette PumpTM (Blue)
Weighing boats
Micropipettors (V-10, V-200, V-1000)
Pipette tips
1.5 ml Microcentrifuge tubes
Deionized water
50 ml beakers
70% Sucrose
Aseptic Techniques Exercise
70% Ethanol
50 ml Beakers
1% Methylene blue
Parafilm
Bunsen burners
Marking Pens
Petri dishes with nutrient agar
Culture tubes with nutrient broth
Inoculating loops
Glass spreading rods
Sterile dilution tubes
Nonsterile dilution tubes
1 and 10 ml Glass pipettes
Pipette PumpTM (Blue and Green)
Escherichia coli cultures
INTRODUCTION
Many genetic laboratory procedures require the transfer of precise amount of liquid from one container to another. You will employ reusable glass pipettes and micropipettes to transfer liquids and bacteria in many of the genetic experiments contained in this manual. Success in these experiments will largely be determined by your ability to properly use these pipettes.
The success of genetic experiments using fungi and bacteria depends upon the use of techniques that prevent the introduction of contaminants into a pure culture. A pure culture contains only one kind of microorganism. Many microbes are present on your workbench, hands, and transfer instruments, as well as in the air. In working with microorganisms, you must transfer growing organisms (called the inoculum) from a pure culture to a sterile medium without introducing any unwanted outside contaminants. This method of preventing unwanted microorganisms from gaining access to your culture is termed aseptic technique.
OBJECTIVES
To learn correct pipetting techniques, aseptic techniques, and how to culture bacteria.
PIPETTING
Using Glass Pipettes
1. In the genetics laboratory, you will never mouth pipette because organisms and/or fluids could accidentally be ingested. All pipetting will be done with a Pipette Pump. Blue Pipette Pumps are for 1-ml pipettes, and green ones are for 5 and 10 ml pipettes.
2. When attaching a Pipette Pump to your pipette, grasp the pipette near its blunt end, touching only the upper portion of the pipette, and press the pipette into the Pipette Pump gently, without excess force.
3. Rotate the wheeled knob with your thumb moving the Pipette Pump plunger up. This should pull the solution up into the pipette. If the Pipette Pump leaks it should be replaced. Any attempt to stop the leak by forcing the pipette further into the Pipette Pump could cause breakage.
4. Once the correct volume has been pulled up into the pipette, place the tip of the pipette against the side of the new container and expel the fluid by rotating the wheeled knob. This should empty the contents of the pipette into the new container. For fast emptying, press down on the plunger top.
Operation of Electronic Balance
The following instructions pertain to the operation of the Ohaus electronic balance:
1. Turning the balance on – Press ReZero On and hold briefly. The balance display should light up and read 0.00. Allow 5 minutes warm-up time.
2. Weighing
a. The electronic balance can weight in the following units: g, dwt, pc, and oz t. Select g (which is indicated in the balance display) by momentarily pressing Mode Off.
b. If the balance does not read 0.00, momentarily press ReZero On to rezero the balance.
c. Place item(s) to be weighed on the pan and read the weight on the balance display. The stability indicator (indicated by an *) to the left of the weight appears when the reading is stable.
3. Taring – When weighing items that must be held in a container, taring subtracts the container’s weight from the total weight on the pan of the balance.
a. With an empty container on the pan, press ReZero On to zero the balance display.
b. As material is added to the container, the new weight is displayed. Tared weight remains in the balance memory until ReZero On is pressed again.
Pipetting Exercise
1. Work in groups of three or four students.
2. Fill a 50 ml beaker with deionized water (density = 1 g/cm3).
3. Attach a blue Pipette Pump to a 1 ml glass pipette.
4. Place an empty weighing boat on an electronic balance and then press ReZero On to zero the display.
5. Pipette 1 ml of deionized water into the weighing boat.
6. Record the weight of the aliquot in your laboratory notebook.
7. Repeat the process 10 times.
8. Each student in the group should perform this pipetting exercise.
9. Data analysis
a. Determine the mean for each set of 10 measurements by adding up the measurements and dividing by 10.
[pic]
where the X with the bar over it (read xbar) is the mean, Xi represents the individual data points, and there are n such points.
b. Determine the standard deviation. The standard deviation expresses the variability of data about the mean. Thus, standard deviation is the average amount that a set of numbers differ from their mean. The closer a set of numbers are to each other, the more consistent they are. Generally, the most consistent set of numbers is usually the one with the lowest standard deviation.
[pic]
c. Determine percentage error. Percentage error is used to represent the relationship of the standard deviation to the mean, telling scientists how much the mean truly represents the numbers it came from. The formula is:
Percentage Error = (Standard Deviation / Mean) * 100
Using a LabnetTM Micropipettor
The micropipettor is a precision instrument designed to measure and transfer extremely small amount of liquids. The scales on micropipettors are in microliters (µl). One microliter is one thousandth of a milliliter (1 µl = 0.001 ml).
You will be using three LabnetTM micropipettors that deliver specified volumes of fluid with the appropriate tip in place. They are:
|Micropipette Top Color |Range of Volumes |Color of Tip to Use |
|White (V-10) |0.5 - 10 µl |White |
|Yellow (V-200) |20 - 200 µl |Yellow |
|Blue (V-1000) |100 - 1000 µl |Blue |
Directions for use:
1. Select the correct micropipettor for the operating range required (See table above).
2. Use the knurled nut (calibration knob) in the handle to set the desired volume. Turn the calibration knob clockwise to increase the volume or counterclockwise to decrease the volume. Never exceed the upper or lower limits of these micropipettors.
3. The display in the handle of micropipettor consists of three vertical numbers. Set the micropipettor to the desired volume by turning the calibration knob until the volume display reads the desired numbers. Examples are given below for each of the three types of micropipettors (V-10, V-200, and V-1000) that will be used in the genetics laboratory.
|V-10 |V-200 |V-1000 |
|(0.5-10.0 µl) |(20-200 µl) |(100-1000 µl) |
| | | |
|0 10's |0 100's |0 1000's |
|1 1's |2 10's |2 100's |
|5 0.1's |5 1's |0 10's |
| | | |
|1.5 µl |25 µl |200 µl |
| | | |
|1 |1 |1 |
|0 |5 |0 |
|0 |0 |0 |
| | | |
|10.0 µl |150 µl |1000 µl |
4. Select the correct tip and place it firmly on the nose of the micropipettor using a slight screwing action. Never use the micropipettor without a tip attached. If sterile conditions are necessary, do not allow the pipette tip to touch any object, including your hands.
5. The plunger of the micropipettor will stop at two different positions when it is depressed. This first of these stopping points is the point of initial resistance and is the level of depression that will result in the desired volume of liquid being transferred. The second stopping point is used to completely discharge liquids from the plastic pipette tip. You should not reach this second point when drawing liquid into the micropipettor, only when expelling the last drop. Before continuing, practice depressing the plunger to each of these stopping points until you can easily distinguish between these points.
6. Prior to immersing the pipette tip in the fluid, depress the plunger to the first stop.
7. Hold the micropipettor vertically and immerse the plastic pipette tip to a depth of 3-5 mm in the fluid.
8. Carefully and slowly release the plunger in a controlled manner to aspirate the selected volume of liquid. If you are pipetting a viscous solution, allow the pipette tip to fill to the final volume before removing it from the solution to avoid the presence of air bubbles in the plastic tip, which will result in an inaccurate volume.
9. Always keep the micropipettor in the vertical position when in use and never lay down a micropipettor with a filled pipette tip.
10. Dispense the liquid into the appropriate container by depressing the plunger to the second resistance stop. Keep the plunger in the depressed position and slide the pipettor out of the container with the plunger depressed. This will avoid the possibility of sucking any liquid back into the tip.
11. Eject the used pipette tip into a disposable container by pressing the ejector button on the micropipettor.
Pipetting Exercise with Micropipettor
1. Obtain a 50 ml beaker and place a small amount of deionized water into it.
11. Pipette the following volumes of water into a 1.5 ml microcentrifuge tube using the correct micropipettors and pipette tips:
a. 350 µl - Set the V-1000 micropipettor (100-1000 µl) fitted with a blue pipette tip to 350 µl [035]. Dispense this volume into the microcentrifuge tube.
b. 90 µl - Set the V-200 micropipettor (20-200 µl) fitted with a yellow pipette tip to 90 µl [090]. Dispense this volume into the microcentrifuge tube.
c. 10 µl - Set the V-10 micropipettor (0.5-10 µl) fitted with a white pipette tip to 10 µl [100]. Dispense this volume into the microcentrifuge tube.
3. When several solutions are added to a microcentrifuge tube, some of the liquid may cling to the walls of the tube. The solutions may be pooled at the bottom of the tube by "pulsing" the tubes in a microcentrifuge. When pooling the contents of a tube, the microcentrifuge tubes are placed in the rotor of the centrifuge in a balanced configuration and the timer is set to zero. Press and release the start button one or twice.
4. A total of 450 µl was pipetted into the microcentrifuge tube. To determine the accuracy of your measurements, set the V-1000 (100-1000 µl) micropipettor to 450 µl and withdraw the contents of the microcentrifuge tube into a blue pipette tip. If you have measured correctly, the entire contents of the microcentrifuge tube should be withdrawn into the pipette tip and there should be no bubbles in the pipette tip or liquid left in the microcentrifuge tube.
5. If you have measured incorrectly, repeat the pipetting exercise again.
ASEPTIC TECHNIQUES
Special care must be taken when handling bacterial and fungal cultures. Aseptic techniques are the procedures for transferring microorganisms from one medium to another without contaminating the cultures or the surrounding environment. A list of general rules for aseptic techniques is given below:
1. Wipe down your work area with 70% alcohol or a bactericidal disinfectant at the beginning and at the end of each laboratory period.
2. Wash your hands before and after handling microorganisms.
3. Coughing, speaking, sneezing or breathing directly over sterile items or cultures can result in contamination. Long hair should be tied back because it is a source of contamination and as a safety precaution around Bunsen burners.
4. Sterilize all inoculation instruments before and after sampling microorganisms.
5. Do not touch parts of a sterile item, which may later come in direct contact with the culture and thus contaminate it.
6. Never lay caps, plugs or sterile items on the bench top. Putting sterile items on the bench and then reusing them results in contamination.
Inoculating Loops
1. Inert metal wires are often used to transfer microorganisms from one culture to another. These loops must be sterilized before being used to transfer microorganisms. To sterilize the loop, grasp its handle and hold the wire in a flame until it is red-hot. Flaming incinerates any organisms on the wire.
2. After cooling, the loop can be used to lift bacteria off surfaces or out of broths and transfer them to other media. The instrument is resterilized after each use.
Pipets
You will employ various pipets and pipettors to transfer liquids and bacteria. The pipettes used in the genetics laboratory are reusable, glass pipettes. They are stored in cans and sterilized. The following precautions should be taken to maintain the sterility of a can full of pipettes:
1. Keep the can closed when not in use.
2. Do not reach over an open can of pipets.
3. Touch only one pipet at a time.
4. To withdraw a pipet, lift it out; do not drag the tip over the blunt ends of other pipets.
5. After obtaining a sterile pipet, use it immediately. NEVER return a pipet to the can, even if you think the pipet is probably sterile.
6. Contaminated (used) pipets should be placed into a container with disinfectant. They are placed into the container gently to avoid breakage and tip down to prevent the formation of aerosols.
Alcohol-Flaming
Instruments that are used to handle samples, such as forceps, loops, spreading rods, may require a rapid sterilization. This can be achieved by dipping an appropriate length of the instrument into 70% or 95% ethanol and passing it through and out of the flame. The alcohol is then allowed to burn off. This technique requires caution because flaming alcohol can drip onto your workbench, igniting any flammable material it contacts. The flaming liquid may stream onto your hand if the instrument is held incorrectly and result in fire damage to you and/or your belongings.
Broth Inoculation
1. Obtain a broth culture of Escherichia coli and a tube of sterile nutrient broth.
2. Gently tap the bottom of the culture tube with your finger 5 or 6 time to suspend the bacterial cells in the broth.
3. Hold the inoculating loop in your dominant hand and the broth culture of E. coli in the other hand.
4. Flame the inoculating loop to redness and allow it to cool approximately 10 seconds.
5. Hold the inoculating loop like a pencil in your dominant hand and curl the little finger of the same hand around the cap of the broth culture of E. coli. Gently remove the cap from the culture tube (Figure 1).
[pic]
Figure 1. Procedure for transferring bacterial to a broth culture.
6. Hold the culture tube at an angle and pass the mouth of the tube through the flame of your Bunsen burner. Always hold culture tubes at an angle to minimize the amount of airborne dust or spores that could fall into them.
7. Carefully dip the small loop at the tip of the inoculating loop into the broth culture of E. coli and remove a loopful of inoculum.
8. Remove the inoculating loop, flame the mouth of the culture tube, replace the cap, and place the culture into the test tube rack.
9. Pick up the sterile broth tube and remove the cap with the little finger of your dominant hand as described previously. Do not set the cap down. Flame the mouth of the culture tube.
10. Inoculate the broth by inserting the tip of the inoculating loop into the broth. Gently shake the loop without touching the sides of the tube with the handle of the loop.
11. Flame the mouth of the culture tube and replace the cap. Return the culture tube to the test tube rack.
12. Resterilize the loop by placing it in the flame of your Bunsen burner. Now you may lay the loop down until it is needed again.
13. Label the inoculated culture tube with your name, laboratory section, the name of the bacteria, and the date.
14. Incubate the culture tubes at 37oC for 48 hours.
15. After 48 hours, examine the tubes for contamination. The broth should be turbid because of bacterial growth.
Streak Plate Technique
The streak plate technique is a procedure in which a bacterial culture is streaked over the surface of agar such that individual cells become separated from one another. Each isolated cell grows into a colony (cluster of cells), which theoretically is the progeny of the original single cell. You will prepare a streak plate using the quadrant method.
1. Obtain a broth culture of Escherichia coli and Petri dish containing sterile nutrient agar.
2. Tap the bottom of the culture tube containing the E. coli five or six times to disperse the cells evenly throughout the broth. Return the culture to the test tube rack.
3. Sterilize an inoculating loop and let it cool for 10 seconds. Aseptically remove a loopful of the broth culture, replace the cap, and put the tube back in the rack. (This should be performed as previously described in preparing a broth inoculation)
4. Lift the lid of a nutrient agar Petri plate with your free hand just enough to insert the inoculating loop, but do not remove it completely. Insert your inoculating loop and touch the surface of the agar medium near one edge of the plate and then make three to four continuous streaks across a small area of the plate (area A, Figure 2). Avoid digging into the agar by keeping the loop horizontal during streaking and applying just enough pressure to allow contact with surface.
Figure 2. Quadrant procedure for preparing a streak plate.
5. Close the lid on the Petri plate and resterilize your loop. Let the loop cool for about 10 seconds.
6. Rotate the Petri plate 90o counterclockwise and then partially open the lid. Make four or five continuous streaks from area A into area B, staying near the edge of the plate as shown in Fig. 2. (You can see the streak marks of the loop in area A).
7. Close the lid on the Petri plate and resterilize your loop. Let the cool for about 10 seconds.
8. Rotate the Petri plate another 90o counterclockwise and then partially open the lid. Make four or five continuous streaks from area B into area C, once again staying near the edge of the plate as shown in Fig. 2.
9. Close the lid on the Petri plate and resterilize your loop. Let the cool for about 10 seconds.
10. Rotate the Petri plate another 90o counterclockwise and then partially open the lid. Spread the bacteria from area C across the rest of the plate as shown in Fig 2.
11. Close the lid on the Petri plate and resterilize your loop. Now you may lay the loop down until it is needed again.
12. Seal the Petri dish with a layer of Parafilm or tape around the edge. This keeps the agar from drying out while it is in the incubator.
13. Invert the Petri plate and label the bottom of the dish with your name, laboratory section, the date, and the name of the organism used to inoculate the culture. Write this information along the outer edge of the plate bottom so that you can observe the culture clearly once it has grown.
14. Invert your streak plate and incubate at 37oC until the next laboratory period.
15. Examine your streak plate for well isolated colonies of E. coli. The plate should be free of contamination.
Spread Plate Technique
If a concentrated solution of bacteria were grown on an agar plate, the colonies would merge and be impossible to count. A method for isolating single colonies of bacteria involves diluting a fraction of a culture such that fewer and fewer cells exist per unit volume of liquid. An aliquot of diluted cells is pipetted onto the surface of an agar plate and the cells are distributed with a glass spreading rod. After incubation, the number of colonies is counted. It is usually possible to obtain accurate counts of 30 to 300 colonies per plate.
Serial Dilution
A bacterial sample is mixed into a series of volumes of sterile diluent that can be used for quantifying the number of bacteria in a sample or for colony isolation
Practice Exercise
1. For practice, you will use a solution of 1% methylene blue that will be serially diluted into a series of 9 ml water blanks.
2. Label dilution tube for the following dilutions: 10-1, 10-2, 10-3, 10-4, 10-5, 10-6
3. Using a 10 ml glass pipette equipped with a Pipette Pump, add 9 ml of deionized water to each tube.
4. Using a 1 ml glass pipette equipped with a Pipette Pump, perform the following serial dilution (1 ml into 9 ml = 1:10 dilution)
a. Add 1 ml of 1% methylene blue to the tube labeled 10-1
b. Add 1 ml of 10-1 solution to the tube labeled 10-2
c. Add 1 ml of 10-2 solution to the tube labeled 10-3
d. Add 1 ml of 10-3 solution to the tube labeled 10-4
e. Add 1 ml of 10-4 solution to the tube labeled 10-5
f. Add 1 ml of 10-5 solution to the tube labeled 10-6
g. Observe the color change through the various dilutions.
Preparation of Spread Plate
1. Label six sterile dilution tube for the following dilutions: 10-1, 10-2, 10-3, 10-4, 10-5,
10-6
2. Using a sterile 10 ml glass pipette equipped with a Pipette Pump, add 9 ml of sterile broth to each tube. You must use aseptic techniques.
3. Obtain six Petri plates containing sterile nutrient agar. Label the plates: 10-1, 10-2, 10-3,
10-4, 10-5, 10-6. Label the bottom of the dish with your name, laboratory section, the date, and the name of the organism used to inoculate the culture. Write this information along the outer edge of the plate bottom so that you can observe the culture clearly once it has grown
4. Obtain a broth culture of E. coli. Tap the bottom of the culture tube containing the E. coli five or six times to disperse the cells evenly throughout the broth. Return the culture to the test tube rack.
5. Prepare serial dilutions as you did in the practice exercise: 10-1 - 10-6. Remember to utilize aseptic techniques.
6. Using a 1 ml sterile glass pipette equipped with a Pipette Pump, transfer 0.1 ml of the 10-1 inoculum onto the agar surface near the center of the appropriate Petri plate. Remember to utilize aseptic techniques.
7. Sterilize a glass spreader by first submersing it in 70% ethanol. Quickly pass the spreader through the flame of a Bunsen burner, and allow the alcohol to burn off. Cool the spreader for 10 seconds. Use the sterile spreader to spread the bacterial solution uniformly over the surface of the plate. Hold the lid over the Petri plate while carrying out this technique to reduce the likelihood of contamination. Do not put the lid down on the bench.
8. Repeat the process for each of your dilutions.
9. Seal each Petri plate with Parafilm or tape.
10. Invert your spread plates and incubate at 37oC until the next laboratory period.
11. Count the number of colonies on each of your Petri plates. If a plate has more than 300 colonies, express the number as too numerous to count. Record the data in your laboratory notebook.
12. Data analysis:
a. Determine the total or final dilution for your cultures.
Example:
Step 1: Add 1 ml sample to 9 ml diluent and mix (1:10)
Step 2: Add 1 ml from step 1 to 9 ml diluent and mix (1:10)
Step 3: Add 1 ml from step 2 to 9 ml diluent and mix (1:10)
Each dilution in this series is a 1:10 dilution or 10-1
Final dilution (product of all dilutions) = 1/10 x 1/10 x 1/10
= 1/1000 (1:1000 or 1 x 10-3)
b. The dilution factor is the reciprocal of the final dilution. For the above example, the dilution factor is 1000 or 103
c. Determine the number of bacteria contained in your sample
Knowing the number of colonies on the plate, the sample dilution plated on the plate, and the amount of the sample dilution plated on the plate, the number of viable bacteria present per milliliter (colony forming units (cfu)/ml) of the original sample can now be determined. This can be accomplished using the following equation:
cfu/ml = Number of colonies x dilution factor x amount of sample plated
For example, if 50 colonies grew when a 0.1 (10-1) ml sample of a 10-4 dilution was plated, the concentration of viable cells in the original culture was:
1 x 104 x 101 = 50 x 105 or 5 x 106 cfu/ml
d. If dilutions were done correctly, there will be a noticeable 10-fold difference in the colony counts of subsequent plates.
EXERCISE 4
LINKAGE, CROSSING OVER, AND GENETIC
MAPPING IN DROSOPHILA MELANOGASTER
EQUIPMENT
Stereo Dissecting Microscopes
SUPPLIES AND MATERIALS
yellow crossveinless vermilion forked (y cv v f) stock of Drosophila melanogaster
wild type Drosophila melanogaster
Instant Medium Formula 4-24® (Carolina Biological Supply Company)
Fly-Nap® (Carolina Biological Supply Company)
Anesthetizing bottles
Camel's hair brushes
Dissecting needles
Morgue
Labeling tape
3 x 5 cards
INTRODUCTION
Sutton (1902) proposed that each chromosome must have more than one gene. Genes that are in the same chromosome are said to be linked and do not follow Mendel's law of independent assortment. If two or more genes are always transmitted together, they are completely linked. However, linked genes are not always kept together in the formation of gametes and hence exhibit incomplete linkage. This is because homologous chromosomes exchange parts during meiosis. The exchange of genes between homologous chromosomes is called crossing over. As a consequence of crossing over, linked genes may be transmitted to progeny in combinations different from those in which they are present in the parents.
Crossing over makes it possible to construct linkage maps. If genes are linearly arranged along chromosomes, the farther apart they are in the chromosome the more likely it is that crossing over will occur between them. Thus, the frequency with which crossing over occurs between linked genes is a function of the distance between them. One map unit is the length of the chromosome within which one percent crossing over occurs. Per cent crossing over is calculated using the formula:
% Crossing over = Total number of recombinants observed
Total number of offspring
Valuable information on linkage relationships can be obtained when the cross involves three genes in the same chromosome. The X chromosome of Drosophila melanogaster is a convenient one for studying not just linkage but more specifically sex linkage. Female fruit flies possess two X chromosomes whereas males have one X chromosome and one Y chromosome. The Y chromosome of the male contains no homologues of the genes in the X chromosome. If a triply recessive female (abc/abc) is crossed with a wild-type male (ABC/Y), the F1 male receives his X chromosome (abc) from his mother and the Y chromosome from his father. The F1 female receives an X chromosome from both parents and will be (ABC/abc). She will produce eight types of gametes (Table 1). Since crossing over does not occur in the male Drosophila, the F1 male will produce only two types of gametes, i.e., abc and Y. It is evident that we can study linkage values if we consider only male progeny. Since males are hemizygous for all genes in the X chromosome, the F2 male phenotypes are exactly like the gametes produced by the F1 female fly (Table 1). Thus, the X chromosome is highly advantageous for studying linkage. It is possible to determine the linkage of genes on the X chromosome without making a testcross.
In a three-point sex-linked cross, the noncrossover (parental) classes will contain the most flies (Table 1). In addition, to the single crossovers between A and B and between B and C there occur also the double crossover classes AbC and aBc. The double crossovers are recognized as the numerically smallest classes and may be used to determine gene order.
In most experiments involving linkage of three genes, it has been found that crossing over at one region of the chromosome interferes with crossing over at other regions. This phenomenon is known as interference. Interference appears to be greatest near centromeres and at the ends (telomeres) of chromosomes. Degrees of interference are commonly expressed as coefficient of coincidence:
Coefficient of coincidence = Actual Number of Double Crossovers
Calculated Number of Double Crossovers
Interference = 1 - Coefficient of coincidence
Coefficient of coincidence values ordinarily vary between 0 and 1. Absence of interference give a coincidence value of 1, whereas complete interference results in a coincidence of 0. In Drosophila, coincidence is 0 for map distanced less than about 10 map units.
Table 1. All Possible Gametes Produced by an F1 Heterozygous
Female (ABC/abc) for Three Sex-linked Genes in Coupling Phase
A B C
Parental types
a b c
A b c
Single Crossover in Region 1
a B C
A B c
Single Crossover in Region 2
a b C
A b C
Double Crossover
a B c
OBJECTIVE
To perform a three-point sex-linked cross, construct a linkage map based on recombination percentages, and determine the amount of interference.
PROCEDURE
1. Obtain a pint milk bottle and add: 3 small measuring cups of instant medium, 3 small cups of 0.5% propionic acid, and add a pinch of yeast to the top of the instant medium. Use a foam stopper to plug the opening of the bottle.
2. Transfer flies from the culture vial into the anesthetizing vial. Insert an anesthetizing wand into the Fly-Nap® and then place the wand in the anesthetizing vial.
3. Once the flies are anesthetized pour them onto a white card and examine them under the dissecting microscope. Be sure that you can tell the mutants apart from wild type flies.
4. Cross 6 virgin females homozygous (y cv v f/y cv v f) for the sex-linked recessive genes yellow crossveinless vermilion forked with 4-6 wild-type males (+ + +/Y). An alternative procedure would be to make the reciprocal cross: +++/+++ virgin females with y cv v f/Y males.
5. Remove the parents once larval activity is visible in the medium (6-7 days). Label this bottle F1. Place the parents on new medium and retain them as a backup in case something happens to the first cross. Label this bottle P1.
6. Examine the F1 and record the phenotypes of the F1 males and females.
7. Set up 3 to 4 P2 crosses by taking 4-6 of the F1 females and crossing them with 4-6 F1 males in each bottle. Label this bottle F1 x F1.
8. Remove the parents once larval activity (6-7 days) is visible in the medium and label the bottle F2. If necessary, place the parents on new medium.
9. Count and score the phenotype of 250 F2 males for a period of 10 days after the first flies begin to emerge. Determine the 8 phenotypic classes before counting and scoring the F2 flies. Since crossing over does not occur in male Drosophila, F2 males will be a direct reflection of the crossover events that occurred during meiosis in the female. Thus, you need count and score only F2 males.
10. The two phenotypic classes with the fewest number of individuals represent double crossover types. The two phenotypic classes with the largest number of individuals represent the parental types.
DATA ANALYSIS
1. Calculate the genetic distance between each pair of genes.
2. Construct a linkage map based on these distances.
3. Calculate the coefficient of coincidence and interference.
SUGGESTED READINGS
Ephrussi, B. and M. C. Weiss. 1969. Hybrid somatic cells. Sci. Amer. 220:26-35.
McKusick, V. A. 1971. The mapping of human chromosomes. Sci. Amer. 224:104-113.
Mertens, T. R. 1972. Investigation of three-point linkage. The Amer. Biol. Teacher 34:523-526.
Offner, S. 1996. A plain English map of the chromosomes of the fruit fly Drosophila melanogaster. The Amer. Biol. Teacher 58:462-469.
Ruddle, F. H. and R. S. Kucherlapati. 1974. Hybrid cells and human genes. Sci. Amer. 231:36-44.
Sutton, W. S. 1902. The chromosomes in heredity. Biol. Bull. 4:231-248.
EXERCISE 5
TETRAD ANALYSIS IN SORDARIA FIMICOLA
EQUIPMENT
Compound Microscope
SUPPLIES AND MATERIALS
Laboratory Session 1
70% Ethanol
Bunsen burners
Wild-type Sordaria with black ascospores
Mutant Sordaria with tan ascospores
Media for crossing Sordaria
Microspatula (sterile)
Marking Pens
Laboratory Session 2
Microscope slides
Cover glasses
Dissecting needles or toothpicks
Water
CULTURE MEDIUM
Corn meal agar 17.0 g
Glucose 2.0 g
Yeast extract 1.0 g
Distilled water 1.0 liter
Suspend the corn meal agar in the distilled water and add the glucose and yeast extract. Plug and sterilize in an autoclave at 15 pounds for 15 minutes. Pour into Petri dishes.
CROSSING MEDIUM
Corn meal agar 17.0 g
Sucrose 10.0 g
Glucose 7.0 g
KH2PO4 0.1 g
Yeast extract 1.0 g
Distilled water 1.0 liter
Suspend the corn meal agar in the water and add the two sugars, salt, and yeast extract. Plug and sterilize at 15 pounds pressure for 15 minutes. Pour into sterile Petri dishes. A well-filled, deep Petri dish should be used to insure adequate moisture for Sordaria growth.
INTRODUCTION
Fungi are convenient organisms for use in genetics because they are haploid, grow rapidly, are easy to manipulate, and require little space. Sordaria fimicola is an ascomycete or sac fungus that has been used extensively in genetics. This species can be found growing in rotting vegetation and animal dung (in fact, the name Sordaria fimicola means "filthy dung dweller"). Sordaria has been used as a model system for studying the process of crossing-over because of its reproductive characteristics.
Sordaria is haploid throughout most of its life cycle and reproduces by sexual spores called ascospores. The conspicuous vegetative stage has a mycelium of branched filaments called hyphae. The hyphae of the mycelium are haploid. When the hyphae of different individuals come in contact (Fig. 1), cells in the hyphae may fuse (plasmogamy) to form a single cell with two nuclei, one derived from each individual. Such a cell is called a dikaryon (literally "double nucleus") or heterokaryon (literally "different nucleus"). The dikaryon is a N+N genotype; it is not actually diploid, because it has two separate nuclei, each of which is haploid. After many mitotic divisions, (forming a large mass of cells, the nuclei in some of the dikaryon cells fuse to form a single diploid nucleus in each cell. Such cells may be properly referred to as zygotes. The zygotes then undergo meiosis to form four haploid nuclei, which are held within a single membranous sac called the ascus (pl. = asci). Many asci are contained within a larger fruiting body called the perithecium. Sordaria forms several hundred perithecia within 5 to 10 days after inoculation on culture agar. In nature, the perithecium eventually ruptures and releases thousands of ascospores into the wind. Thus, the life cycle is completed. As you can see, the great majority of the life cycle of this species is haploid. In some ascomycetes, the ascospores are arranged randomly within the ascus, but in Sordaria, they are arranged in a way that reflects the events that took place during the first and second meiotic divisions. The four products of meiosis (in any organism) are called a tetrad. When an additional mitotic division occurs, as in ascomycetes, the tetrad becomes an octad. However, since we are really only interested in the products of the meiotic events, we generally view the ascospores as four pairs. Hence, we perform tetrad analysis (rather than "octad analysis").
[pic]
[pic][pic]
Figure 1. Life cycle of Sordaria.
In this experiment we will be concerned with the phenotype of the spores produced in a cross of a Sordaria strain having wild-type black ascospores (+) with a mutant strain that has tan-colored spores (t). The sequence of spores in the ascus reflects the products of the first and second meiotic divisions. Since the spores of an ascus are contained in the linear order in which they were produced during meiosis, it is possible to visualize the meiotic events that must have occurred to produce any particular spore order under consideration. Some asci will have the spore arrangement ++++ t t t t or t t t t ++++ where wild type is represented by + and tan by t (Fig. 2). These spore arrangements represent a first-division segregation for the spore color gene and indicate that a crossover did not occur between the gene and its respective centromere during meiosis. A second-division segregation reflects a crossover between the gene and its centromere and produces a variety of spore patterns (e.g. ++ t t ++ t t; t t ++ t t ++; ++ t t t t ++; t t ++++ t t).
In this exercise you will undertake an ordered tetrad analysis of the Sordaria cross. The analysis of ordered tetrads allows for the mapping of centromeres, since the
occurrence of second division segregation is revealed by the arrangement of the ascospores in the ascus. The closer a gene is to its centromere, the fewer times a crossover will occur between that gene and its centromere; conversely, the farther apart the gene and its centromere, the more frequently crossing over will occur. Thus, the distance separating a gene from its centromere is reflected by the frequency of recombinants found in the progeny.
OBJECTIVE
To introduce you to genetic analysis using a haploid eukaryotic organism and to map the distance between a gene and its centromere using ordered tetrad analysis.
[pic]
Figure 2. Illustration of ascospore arrangement after first and second division segregation. + = black or wild type; t = tan.
Laboratory Session 1
CROSSING PROCEDURE – WEEK ONE
1. Obtain a Petri dish containing crossing medium. Invert the Petri dish and with a marking pen divide the bottom of the dish into four sections (Fig. 3).
Figure 3. Procedure for positioning Sordaria on crossing medium.
2. Sterilize a narrow stainless steel microspatula by dipping it in ethanol and flaming it with a Bunsen burner. Sterile techniques must be used in performing this cross or the cross will become contaminated and be useless for tetrad analysis.
3. Before obtaining Sordaria, check the temperature of the microspatula by touching it to the agar. If the agar sizzles, wait to obtain Sordaria until the microspatula cools.
4. Using your sterilized microspatula, remove a ~ 5 mm square block of mycelium (with no perithecia) from the culture containing the wild (black) Sordaria. Place the agar plug upside down on the crossing medium in one of the sections marked black (Fig. 3). Never remove the top of the Petri dish. Open the top of the Petri dish just enough to insert the sterile microspatula and obtain a sample.
5. Resterilize the microspatula and repeat step 4. Place the black Sordaria mycelium on the second section of the Petri dish marked black about 2-3 cm from the previous sample.
6. Repeat steps 2-5 using the tan ascospore mutant. Place the tan mutant immediately adjacent to the wild-type mycelium (Fig. 3).
7. Put your name, laboratory section, and date on the Petri dish and place it in the dark. Observe the plates throughout the next seven days, as you keep them at room temperature and out of direct sunlight. Make notes in your laboratory notebook on the appearance of the plates. During this time the hyphae from each mycelial block are growing out and toward hyphae from the other blocks. Where they meet they will fuse and form mating structures culminating in black dots, the perithecia. So you should see on your plates a rough “X” of black dots. These cultures will mature in 7-10 days and should contain visible perithecia.
8. After a week remove a few perithecia with a sterilized needle and place them in a drop of water on a slide. Cover the perithecia with a coverslip. If tapping on the coverslip with a pencil eraser or gentle pressure with your thumb easily breaks the perithecia open and releases lots of asci with mature (observable) ascospores, the cross is ready to score. If tapping does not break open the perithecia, or if there are only a few asci released which contain underdeveloped ascospores, allow the crosses to mature for another day.
9. When the perithecia are mature, continue with the experiment. If it is inconvenient to continue when the perithecia are mature, refrigerate (4°C) the cross plate until you are ready to continue the experiment.
Laboratory Session 2
COLLECTING THE DATA
1. Use a toothpick or dissecting needle to remove a few dark perithecia from the crossing dish. If the cross was successful, you should be able to see where the mycelia from the two strains meet in the center of the Petri dish. Perithecia taken from this area should contain hybrid ascospores.
2. Place the perithecia in a drop of water on a microscope slide and add a coverglass.
3. Gently press on the coverglass to crush the perithecia and reveal the asci with ascospores. The job now is to gently rupture the perithecia to release the asci, but to not rupture the asci in the process. Students have a tendency to either squash the perithecium so hard the asci rupture or too little so that the asci are not released from the perithecium. You may need to do this step several times to get a good preparation: lots of intact asci visible under the microscope.
4. Using the low power objective of a microscope, search for hybrid asci (asci that contain both black and tan ascospores). Do not count asci with only tan or only black ascospores
5. After locating hybrid asci, examine the preparation, using 400x magnification. If the preparation is successful, the asci will be spread out in a "sunburst" arrangement. Begin at the "12 o'clock position" and score each ascus possible, moving in a clockwise direction. Score at least 100 asci as to whether each contains a first division segregation or a second division segregation.
6. When wild-type Sordaria are crossed with the tan mutant, hybrid asci are produced that contain four dark (wild-type) and four light (mutant) ascospores. The order of ascospores in the ascus reflects the order in which the chromosomes are segregated during meiosis. If crossing over has occurred, the sequence of mutant to wild-type spores will be 2:2:2:2 or 2:4:2. If crossing over has not occurred, the sequence will be 4:4. The 2+:2t:2+:2t arrangement is the result of crossing over between chromatids two and three, whereas if chromatids one a four crossover a 2t:2+:2t:2+ pattern results. The 2+:4t:2+ pattern results from crossing over involving chromatids two and four, whereas crossing over between chromatids one and three gives the 2t:4+:2t pattern.
7. Map units between the gene and its centromere can be calculated using the following formula:
Map units = 1/2(# of asci showing 2nd division segregation) x 100
Total Number of Asci
DATA ANALYSIS
Number of asci scored _______________________________________
Number of second division segregations__________________________
Percent second division segregation _____________________________
Genetic map units between tan locus and its centromere______________
SUGGESTED READINGS
Bistis, G. and L. S. Oliver. 1954. Ascomycete spore mutants and their use in genetic studies. Science 120:105-106.
Carr, A. J. H. and L. S. Oliver. 1958. Genetics of Sordaria fimicola. II. Cytology. Amer. J. Bot. 45:142-150.
Cassell, P. and T. R. Mertens. 1968. A laboratory exercise on the genetics of ascospore color in Sordaria fimicola. The Amer. Biol. Teacher 30:367-372.
El-Ani, A. S., L. S. Olive and Y. Kitani. 1961. Genetics of Sordaria fimicola. IV. Linkage group I. Amer. J. Bot. 48:716-723.
Ellis, C.H. 2000. Neurospora and tetrad analysis.
Oliver, L. S. 1956. Genetics of Sordaria fimicola. I. Ascospore color mutants. Amer. J. Bot. 43:97-107.
EXERCISE 6
BIOCHEMICAL GENETICS: CHROMATOGRAPHIC
ANALYSIS OF DROSOPHILA EYE PIGMENTS
EQUIPMENT
Chromatography Chamber (1/group)
UV light
UV Goggles
SUPPLIES AND MATERIALS
Chromatographic filter paper (8 in x 8 in)
Ruler (1/group)
Glass rod (1/group)
Razor blade or scalpel (1/group)
Etherizer
Fly-Nap® (Carolina Biological Supply Company)
28% Ammonium hydroxide
n-Propanol
CHROMATOGRAPHIC SOLVENT
Fill the chromatography jars with 50 ml of solvent (1:1 mixture of 28% NH4OH and n-propanol) before the start of the experiment. This will allow for equilibration of solvent in the jar.
DROSOPHILA EYE MUTANTS
Wild (+)
White (w)--affects pigment deposition in eye
Brown (bw)--affects pteridine synthesis
Sepia (se)--affects pteridine synthesis
Cinnabar (cn)--affects ommochrome synthesis
Scarlet (st)--affects ommochrome synthesis
Vermilion (v)--affects ommochrome synthesis
INTRODUCTION
Chemical reactions are constantly occurring within cells, and each reaction is catalyzed by a specific enzyme. Thus, the metabolism of an organism consists of a series of interrelated enzymatic reactions. The study of metabolic pathways in Neurospora crassa by Beadle and Tatum lead them to formulate the one gene-one enzyme hypothesis. This hypothesis proposed that metabolic events are mediated by enzymes under genetic control. Thus, every biochemical pathway can be resolved into a series of individual steps, each mediated by a different enzyme that is the product of a single gene. A mutation in an enzyme-producing gene is likely to induce a metabolic block in one step of a biochemical pathway. The consequences stemming from an excess of precursors or lack of end product(s) depends on the pathway involved.
We now know that an enzyme may be composed of several polypeptide chains so that, the Beadle and Tatum hypothesis has been restated as the one gene-one polypeptide chain hypothesis. Also, genes code for proteins in general, not just enzymes. Hence, one gene codes for the amino acid sequence of one polypeptide chain. Not all genes code for proteins. Some genes code for RNAs such as transfer and ribosomal RNAs.
The dull red compound eye of Drosophila melanogaster contains two types of pigments, the bright red pteridines and the brown ommochromes. The pteridines are light sensitive and play a role in photoreception. The ommochromes occur in cells separating the photoreceptor cells and prevent reflection of light from one facet of the compound eye to another. The synthesis of these two different eye pigments involves two entirely different biochemical pathways (Figs. 1 and 2). Mutant genes are known which block the synthesis of the ommochromes but have no effect on the pteridines. Such mutant Drosophila are characterized by compound eyes, which have a bright red color. Likewise, mutations in the pathway for pteridine biosynthesis have been found. These Drosophila are characterized by brown eyes.
Chromatography can be used to separate and identify compounds such as Drosophila eye pigments. Slightly different solubilities of compounds in a solvent permit their separation. In paper chromatography, samples are applied to a strip of Whatman filter paper. The filter paper is then placed into a chromatography jar containing the proper solvent. The paper acts as a wick and the compounds will migrate up the paper at different rates. Factors affecting the separation of compounds along the filter paper are a combination of absorption, ion exchange, and partition. The distance traveled by any compound is dependent on the rate of migration of molecules along a moving solvent front produced by the rise of the solvent along the length of the filter paper. Since the more soluble molecules will travel the greatest distance, the relative distance can be measured and the rate of migration can be calculated.
Rf = Distance substance traveled
Distance solvent traveled
[pic]
Figure 1. Biochemical pathway for the synthesis of ommochrome pigments. Mutation in the pathway are shown in parentheses.
Figure 2. Biochemical pathway for the synthesis of pteridine pigments. Mutation in the pathway are shown in parentheses.
OBJECTIVE
To use chromatography to characterize the pigments of various Drosophila melanogaster eye mutants.
PROCEDURE
1. Obtain a 8 inch x 8 inch sheet of chromatography paper. Handle chromatography paper only by the edges and place the paper on top of a clean sheet of notebook paper or a paper towel. Foreign matter, especially fingerprints, on the chromatography paper will affect the results of the experiment.
2. With a pencil, draw a light line across the filter paper parallel to, and about 3/4 inch from the bottom.
3. Place a small "x" at 1-inch intervals along this line. No x's should be closer than 1-inch to either of the side edges.
4. Beneath each "x" write the symbol for one of the Drosophila eye mutants. In the upper right-hand corner write your initials and the sex of the flies you are using.
5. Anesthetize the flies. Sort out five flies of proper sex for each mutant. Because sex differences exist for pteridines, each group should use flies of the same sex. Choose either males or females but do not use both on same chromatogram.
6. Using a razor blade or scalpel, cut off the head of each of the five flies on an index card. Use a dissecting scope if you have difficulty deciding where to cut.
7. Use the white-eyed flies first to prevent contamination and the wild-type flies last.
8. Using a clean glass rod, touch the rod to one head (the head will temporarily stick to the rod) and place the head on chromatography paper on the appropriate "x".
9. Using your glass rod, thoroughly crush one head at a time, making as small a dot as possible.
10. Allow the spot to dry before addition of another head.
11. Dip your razor blade or scalpel and glass rod in water and wipe dry for the next set of three fly heads. Clean glass rod after applying each different eye mutant to avoid contamination of pteridines from various mutants.
12. After all heads have been crushed on the chromatography paper, roll the chromatogram (spots on inside) into a cylinder. Connect the two sides with chromatography clips; be sure the edges do not overlap or touch each other.
13. Place this cylinder into the chromatography chamber, base line down. (NOTE: Solvent depth should be lower than spots on chromatogram). Place aluminum foil around chromatography chamber to protect pteridines, which are light sensitive, from the light.
14. When solvent reaches to about 1/2 inch from top of paper, remove the chromatogram from the chamber and mark the solvent front with a pencil.
15. Stand chromatogram on a piece of paper toweling for approximately 10 minutes to air dry.
16. When the chromatogram is dry, remove clips and flatten. Examine the chromatogram under UV light at 350 nm. Wear goggles when examining chromatograms. DO NOT LOOK DIRECTLY AT THE UV LIGHT. If chromatogram must be kept for more than one day before reading, place it in a cool dry, dark place.
ANALYSIS OF CHROMATOGRAM
1. Visible colors (i.e. detectable without UV light)
2. Using a pencil, circle and label each spot for each Drosophila eye mutant.
3. Visible colors include (in sequence from the bottom of the chromatogram):
|Spot |Color |Chemical |
| 1 |Brown |Ommochrome |
| 2 |Orange-red |Drosopterin |
| 3 |Yellow |Sepiapterin |
4. Colors seen under UV light are (in order from the bottom of the chromatogram):
|Spot |Color |Chemical |
|2 |Orange-red |Drosopterin |
|3 |Deep blue |Isoxanthopterin |
|4 |Green-blue |Xanthopterin |
| 5 |Yellow |Sepiapterin |
|6 |Light blue |2-amino-4-hydroxypteridine |
| 7 |Light blue |Biopterin |
|8 |Yellow |Isosepiapterin |
5. The ommochrome and drosopterin (spots 1 and 2) often occupy the same location. The amount of 2-amino-4-hydroxypteridine in wild-type flies may be small or not detectable. Biopterin, 2-amino-4-hydroxypterin, and sepiapterin may overlap and be hard to distinguish.
6. Computation of Rf (reference front) value for each pteridine spot:
a. Find the approximate center of each spot and mark it with a dot.
b. Measure the distance from the base line to the center of each pteridine spot.
c. Measure the distance from the base line to the solvent front.
d. Rf = Distance of pteridine
Distance to solvent front
Pour your agarose gel for EXERCISE 7: PROTEIN ELECTROPHORESIS AND HEMOGLOBIN POLYMORPHISM. Wrap the gel in plastic wrap and store in refrigerator until the next lab period.
SUGGESTED READINGS
Gregg, T. G. and L. A. Smucker. 1965. Pteridines and gene homologies in the eye color mutants of Drosophila hydei and Drosophila melanogaster. Genetics 52:1023-1034.
Hadorn, Ernest. 1962. Fractionating the fruitfly. Sci. Amer. 206:100-110.
Hanly, E . W. and W. H. Hemmert. 1967. Morphology and development of the Drosophila eye. J. Embryol. Exptl. Morph. 17:501-511.
Reaume, A. G., D. A. Knecht, and A. Chovnick. 1991. The rosy locus in Drosophila melanogaster: Xanthine dehydrogenase and eye pigments. Genetics 129:1099-1109.
Wilson, T. G. and K. B. Jacobson. 1977. Isolation and characterization of pteridines from heads of Drosophila melanogaster by a modified thin-layer chromatography procedure. Biochem. Genetics 15:307-319.
Ziegler, I. 1961. Genetic aspects of ommochrome and pterin pigments. Adv. Genetics 10:101-110, 349-403.
EXERCISE 7
PROTEIN ELECTROPHORESIS AND HEMOGLOBIN POLYMORPHISM
EQUIPMENT
BioRad Electrophoresis cells
Power supply units
Gel Trays (1/group)
Mini-Gel Caster (1/group)
Sample well-forming comb (1/group)
Adjustable micropipettors with tips
Balances
Microwave oven
SUPPLIES AND MATERIALS
Laboratory Session 1
Agarose
Tris-glycine buffer, pH 9.2 or Hemoglobin electrophoresis buffer
100 ml Erlenmeyer flask (1/group)
100 ml graduated cylinder (1/group)
50 ml beaker of distilled water
Plastic Wrap
Kimwipes
Bromophenol blue
Bromocresol purple
Phenol red
Orange G
Wire test tube racks (1/group)
Laboratory Session 2
Agarose
Tris-glycine buffer, pH 9.2 or Hemoglobin electrophoresis buffer
100 ml Erlenmeyer flask (1/group)
100 ml graduated cylinder (1/group)
50 ml beaker of distilled water
Plastic Wrap
Kimwipes
Loading dye
Normal human hemoglobin
Sickle trait hemoglobin
Sickle cell hemoglobin
Staining trays (1/group)
Ponceau S
De-staining solution
Wire test tube rack (1/group)
Loading dye
PONCEAU S
Ponceau S 2.0 gm
Trichloroacetic acid 30.0 gm
Distilled water 1000.0 ml
DYE MIXTURE
Dye (Bromophenol blue, Bromocresol
purple, Phenol red, or Orange G) 0.02 g
Sucrose 5.0 g
Water to 10.0 ml
Dispense each different dye into 1.5 ml Eppendorf tubes.
DE-STAINING SOLUTION
Methanol 200.0 ml
Glacial acetic acid 200.0 ml
Water 1600.0 ml
Store tightly capped at room temperature
0.4 M TRIS-GLYCINE BUFFER, pH 9.2
Tris 207.3 g
Glycine 31.5 g
Water to 6 liters
LOADING DYE
Bromophenol blue 0.025 gm
Xylene cyanol 0.025 gm
Sucrose 5.0 gm
Distilled water to 10.0 ml
INTRODUCTION
Electrophoresis is based on the fact that charged molecules in solution, mainly proteins and nucleic acids, will migrate in an electrical field. Electrophoresis entails placing the sample to be analyzed on the end of a gel prepared of agarose, starch, or polyacrylamide and a suitable buffer system. An electrical current is passed through the gel and under its influence the molecules present in the sample move through the gel matrix. The rate at which a molecule moves through the gel depends upon the temperature and pH of the gel, the concentration of the gel matrix, and the size, shape and electric charge of the molecule. After a time, the current is stopped, and the gel is stained. After staining, a series of bands appear in the gel at the position where the molecules have moved.
Electrophoresis has proven to be an important genetic tool. Changes in the amino acid composition of a protein can often be detected by electrophoresis. If an amino acid substitution causes a change in the net electrical charge of the protein, then its mobility in an electric field will be altered. Thus, normal and mutant forms of the protein may migrate at different speeds during electrophoresis.
Hemoglobin variants have been identified in human populations using electrophoresis. Normal adult hemoglobin (hemoglobin A) has four polypeptide chains, two α and β chains. The α and β chains are products of different genes. The α chains contains 141 amino acids and the β chain is composed of 146 amino acids. Each polypeptide chain is attached to an iron-containing heme group. Oxygen binds reversibly to the iron atom in the heme group. One well-known hemoglobin variant is the mutation giving rise to sickle-cell anemia. The sickle-cell mutation is in the gene that codes for the β chain. Sickle-cell hemoglobin (HbS) differs from hemoglobin A (HbA) by a single amino acid substitution of valine for glutamic acid at position six of the β chain. People homozygous for HbS suffer from severe anemia and few of them live to puberty unless given blood transfusions.
Sickle-cell anemia is caused by homozygosity for a mutation in the gene for the ß polypeptide of hemoglobin. The sickle-cell mutation ßS is codominant with the wild-type allele βA. People who are heterozygous (βAβS) make two types of hemoglobin, one called HbA is completely normal with two normal a and two normal β chains. The other, called HbS, is a defective hemoglobin with two normal a and two abnormal β chains specified by the mutant βS gene. Heterozygous individuals (βAβS) suffer from a mild form of sickle-cell anemia called sickle-cell trait.
Hemoglobin A is only one type of hemoglobin found in humans. Different stages of human development are characterized by different hemoglobins. Fetal hemoglobin (HbF) occurs in the blood of the human fetus. It is similar to HbA but has two α and two γ chains. The γ chain is coded for by a gene distinct from the α- and ß-genes. Fetal hemoglobin is produced until just before birth at which time synthesis of the γ chain stops and production of the β chain starts.
OBJECTIVE
This first portion of this exercise is intended to acquaint you with the principle of electrophoresis. In the second portion of the exercise, you will examine the electrophoretic mobilities of normal and mutant hemoglobins and learn how electrophoresis can be used in genetic analyses.
ELECTROPHORESIS PROCEDURE
Laboratory Session 1
Pouring the Agarose Gel
1. Place the Mini-Gel Caster on a level surface. Disengage and slide the movable wall to the open of the caster by turning and lifting the cam peg upward.
2. Place the open end of the gel tray against the fixed wall of the gel caster. Slide the movable wall against the edge of the gel tray and engage the cam peg by turning and pressing downward simultaneously. When the cam peg has dropped into place, turn the peg until resistance is felt. This will seal the edges of the tray for gel casting.
3. Insert the comb into the second slot from the end of the tray.
4. Make a 0.8% agarose gel solution. Place 30 ml of electrophoretic buffer into a 100 ml Erlenmeyer flask or a 100 ml bottle and add 0.24 g of agarose. Gently swirl the flask until the agarose forms a suspension.
5. Heat the agarose in the microwave oven, using 50% power for 30-45 seconds. Be sure to cover flask with plastic wrap or to use a bottle with a loosened cap (to release pressure during heating). Stir frequently during microwaving and stop if the solution starts to boil. The agarose solution should be absolutely clear with no un-melted agarose visible. The correct protocol for microwaving the agarose is to mix the agarose in the buffer, microwave, stop, mix, microwave, etc.
6. Allow the agarose solution to cool to 50oC (check with thermometer). Hot agarose may cause the gel tray and comb to warp or crack.
7. Pour the agarose onto the gel tray until the comb teeth are submerged in gel solution (~20 ml). Remove any air bubbles from the agarose.
8. Allow the gel to cool for 15 minutes. The gel must be completely solidified before the comb is removed or the wells may be improperly formed and the hemoglobin bands distorted.
9. Carefully remove the comb from the solidified gel by lifting it straight out of the gel slowly.
10. Disengage the cam peg by turning and lifting upward. When the gel is solidifying, a light seal is formed between the gasket and the gel. Before moving the wall of the gel caster, carefully lift the gel tray on one side to release the seal or use a spatula to break the seal between the agarose and gasket. Gels can be stored for one week before use. For gel storage, the comb is left in place and the tray containing the gel and comb is wrapped in plastic wrap or placed in a zip lock bag and stored in the refrigerator.
11. Place the gel tray onto the base of the electrophoretic cell so that the sample wells are near the cathode (negative pole or black). The hemoglobin molecules migrate toward the anode (positive pole or red) during electrophoresis.
12. Pour ~275 ml of 1x electrophoretic buffer (Tris-Glycine, pH 9.2) into the buffer tray. The gel should be completely submerged with the buffer 2 to 6 mm above the gel surface.
Loading the Samples
1. Set the V-10 digital micropipettor (uses clear tips) to 10 µl. Place a pipet tip on the micropipettor.
2. Load 10 µl of each of the four colored dyes into a separate sample well as indicated below. Use a different pipet tip for each sample.
3. To load a sample:
a. Use two hands to keep the micropipettor steady over the gel.
b. Expel any air in the pipet before loading the sample.
c. Position the pipette tip under the surface of the buffer and over the well.
d. Slowly expel the sample into the well.
e. Be careful not to punch the pipet tip through the bottom of the well.
|Sample Well Number |Dye |
|1 |Bromophenol blue |
|2 |Phenol red |
|3 |Orange G |
|4 |Bromocresol purple |
|5 |Bromophenol blue |
|6 |Phenol red |
|7 |Orange G |
|8 |Bromocresol purple |
4. Place the lid on the electrophoretic cell carefully so as not to disturb the samples. The lid attaches to the cell in only one orientation. To attach the lid correctly, match the red and black banana jacks on the lid with the red and black banana plugs of the cell.
Electrophoresis
1. Connect the electrophoresis cell to the power supply by inserting the molded two-prong plugs into the power supply’s high voltage output jacks.
2. Press the Power Switch that is located on the side of the power supply. Press the side labeled “I” on the switch.
3. Press the Constant (Const) key to select constant voltage.
4. Use the Scroll Key to enter 100 volts.
5. Press the Run Key to start the run.
6. Run the samples for ~45 minutes.
7. Stop the run by pressing the Run Key.
8. Press the Power Switch to turn the power supply off, disconnect the power cords from the cell, remove the top of the electrophoresis cell, and remove the gel tray containing the gel.
9. Empty the buffer from the electrophoresis cell and store in the refrigerator. Rinse the electrophoresis cell and lid with water.
Analyzing the Gel
1. Carefully slide the gel from the gel tray onto a piece of plastic warp. Thoroughly wash the gel tray with water.
2. Measure the distance traveled by the dyes. Measurements are typically done from the leading edge of the sample well to the leading edge of the dye band.
Pour another agarose gel, wrap the gel in plastic wrap, and store the gel in the refrigerator for use in Laboratory Session 2.
Laboratory Session 2
PROCEDURE
Pouring the Agarose Gel
1. Place the Mini-Gel Caster on a level surface. Disengage and slide the movable wall to the open of the caster by turning and lifting the cam peg upward.
2. Place the open end of the gel tray against the fixed wall of the gel caster. Slide the movable wall against the edge of the gel tray and engage the cam peg by turning and pressing downward simultaneously. When the cam peg has dropped into place, turn the peg until resistance is felt. This will seal the edges of the tray for gel casting.
3. Insert the comb into the slot at the end of the gel tray.
4. Make a 0.8% agarose gel solution. Place 30 ml of electrophoretic buffer into a 100 ml Erlenmeyer flask or a 100 ml bottle and add 0.24 g of agarose. Gently swirl the flask until the agarose forms a suspension.
5. Heat the agarose in the microwave oven, using 50% power for 30-45 seconds. Be sure to cover flask with plastic wrap or to use a bottle with a loosened cap (to release pressure during heating). Stir frequently during microwaving and stop if the solution starts to boil. The agarose solution should be absolutely clear with no un-melted agarose visible. The correct protocol for microwaving the agarose is to mix the agarose in the buffer, microwave, stop, mix, microwave, etc.
6. Allow the agarose solution to cool to 50oC (check with thermometer). Hot agarose may cause the gel tray and comb to warp or crack.
7. Pour the agarose onto the gel tray until the comb teeth are submerged in gel solution (~20 ml). Remove any air bubbles from the agarose.
8. Allow the gel to cool for 15 minutes. The gel must be completely solidified before the comb is removed or the wells may be improperly formed and the hemoglobin bands distorted.
9. Carefully remove the comb from the solidified gel by lifting it straight out of the gel slowly.
10. Disengage the cam peg by turning and lifting upward. When the gel is solidifying, a light seal is formed between the gasket and the gel. Before moving the wall of the gel caster, carefully lift the gel tray on one side to release the seal or use a spatula to break the seal between the agarose and gasket. Gels can be stored for one week before use. For gel storage, the comb is left in place and the tray containing the gel and comb is wrapped in plastic wrap or placed in a zip lock bag and stored in the refrigerator.
11. Place the gel tray onto the base of the electrophoretic cell so that the sample wells are near the cathode (negative pole or black). The hemoglobin molecules migrate toward the anode (positive pole or red) during electrophoresis.
12. Pour 1x electrophoretic buffer (Tris-Glycine, pH 9.2) into the buffer tray until the gel is completely submerged and the buffer is 2 to 6 mm above the gel surface.
Loading the Samples
1. Set the V-10 digital micropipettor (uses clear tips) to 10 µl. Place a pipet tip on the micropipettor.
2. Place 10 µl of each hemoglobin sample onto a small piece of Parafilm. Add 2 µl of loading dye to each hemoglobin sample and mix with the pipet tip. Use a different pipet tip for each sample.
3. To load a sample: A. Use two hands to keep the micropipettor steady over the gel. B. Expel any air in the pipet before loading the sample. C. Position the pipet tip under the surface of the buffer and over the well. D. Slowly expel the sample into the well.
4. Be careful not to punch the pipet tip through the bottom of the well.
5. Load 10 µl of each hemoglobin samples into the sample well as indicated below.
|Sample Well Number |Protein Sample |
|1 |Normal hemoglobin (AA) |
|2 |Sickle trait hemoglobin (AS) |
|3 |Sickle cell hemoglobin (SS) |
|4 |Leave empty |
|5 |Normal hemoglobin (AA) |
|6 |Sickle trait hemoglobin (AS) |
|7 |Sickle cell hemoglobin (SS) |
6. Place the lid on the electrophoretic cell carefully so as not to disturb the samples. The lid attaches to the cell in only one orientation. To attach the lid correctly, match the red and black banana jacks on the lid with the red and black banana plugs of the cell.
Electrophoresis
1. Connect the electrophoresis cell to the power supply by inserting the molded two-prong plugs into the power supply’s high voltage output jacks.
2. Press the Power Switch that is located on the side of the power supply. Press the side labeled “I” on the switch.
3. Press the Constant (Const) key to select constant voltage.
4. Use the Scroll Key to enter 120 volts.
5. Press the Run Key to start the run.
6. Electrophorese the samples for ~45 minutes.
7. Stop the run by pressing the Run Key.
8. Press the Power Switch to turn the power supply off, disconnect the power cords from the cell, remove the top of the electrophoresis cell, and remove the gel tray containing the gel.
9. Pour the electrophoretic buffer back into its original container and store it in the refrigerator. Rinse the electrophoresis cell and lid with water.
Staining and Destaining
1. Gloves should be worn to avoid contact with the staining and destaining solutions.
2. Carefully slide the agarose gel out of the gel tray and place it in a staining dish. Agarose gels are fragile and will break. For this reason, they should be handled with special care. Thoroughly wash the gel tray with water.
3. Cover the gel with Ponceau S (staining solution). Make certain the gel is covered by the stain and does not stick to the staining tray.
4. Stain for 1 minutes, decant and discard the stain, rinse the gel with water, and cover the gel with destaining solution.
5. Change the destaining solution after at least 1 day.
Analyzing the Gel
1. When the background of the gel has destained sufficiently to see the hemoglobin bands on the gel, carefully slide the gel from the staining tray onto a piece of plastic warp.
2. Place the gel onto a light box and note the position of the stained protein bands.
3. Measure the distance (mm) traveled by the hemoglobin. Measurements are typically done from the leading edge of the sample well to the leading edge of the dye band.
4. The gels can be stored in a small plastic bag with a few milliliters of destaining solution. Alternatively, place the stained gel on a dry glass slide and smooth with a gloved index finger to remove air bubbles between the gel and slide. Allow the gel to dry onto the slide for 3-4 days. Cover the dry gel film and glass slide with plastic wrap.
SUGGESTED READINGS
Allison, A. C. 1959. Metabolic polymorphisms in mammals and their bearing on problems of biochemical genetics. Amer. Nat. 93:5-16.
Bank, A., J. G. Mears, and F. Ramirez. 1980. Disorders of human hemoglobin. Science 207:486.
Cerami, A. and C. M. Peterson. 1975. Cyanate and sickle cell disease. Sci. Amer. 232:44.
Doolittle, R. F. 1985. Proteins. Sci. Amer. 253:88.
Ingram, V. M. 1956. A specific chemical difference between the globins of normal human and sickle-cell anaemia haemoglobin. Nature 178:792.
Little, P. F. R., R. A. Flavell, J. M. Kooter, G. Annison, and R. Williamson. 1979. Structure of the human fetal globin gene locus. Nature 278:227-231.
Maniatis, T., E. F. Fritsch, J. Lauer, and R. M. Lawn. 1980. The molecular genetics of human hemoglobins. Ann. Rev. Genet. 14:145-178.
Neel, J. V. 1949. The inheritance of sickle-cell anemia. Science 110:64.
Pauling, L., H. A. Itano, S. J. Singer, I. C. Wells. 1949. Sickle-cell anemia, a molecular disease. Science 110:543.
Perutz, M. F. 1964. The hemoglobin molecule. Sci. Amer. 211:64-76.
Weatherall, D. J. and J. B. Clegg. 1979. Recent developments in the molecular genetics of human hemoglobin. Cell 16:467-479.
EXERCISE 8
RESTRICTION ENDONUCLEASE DIGESTION AND ANALYSIS OF LAMBDA DNA
EQUIPMENT
BioRad Electrophoretic Unit
Power Supply
Gel casting trays (1/group)
Sample well-forming combs (1/group)
Water bath, 37oC
Balances
Adjustable Micropipettors with tips
Photodocumentation unit
SUPPLIES AND MATERIALS
Laboratory Session 1
EcoRI (Restriction enzyme)
Hind III (Restriction enzyme)
Pst I (Restriction enzyme)
2x Restriction Buffer
Lambda DNA (0.4 µg/µl)
Ice bath
1.5 ml Eppendorf tubes (4/group)
Waterproof Marking Pens (1/group)
Loading dye
Laboratory Session 2
InstaStainTM ethidium bromide card or InstaStainTM methylene blue card
Agarose
100 ml Erlenmeyer flask (1/group)
Staining tray
TAE or TBE Buffer
Loading Dye
50X TAE BUFFER
Tris 242.0 g
Glacial acetic acid 57.1 ml
500 mM EDTA 100.0 ml
Dissolve Tris in 600 ml water. Add acetic acid and EDTA and then add distilled water to make 1 liter
500 mM EDTA
Disodium EDTA.2H20 93.05 g
Distilled water 300.0 ml
Stir using a magnetic stirrer. Adjust to pH 8.0 with NaOH (approximately 10 g). Bring volume to 1 liter with distilled water. Dispense into bottle and autoclave.
LOADING DYE
Bromophenol blue 0.025 gm
Xylene cyanol 0.025 gm
Sucrose 5.0 gm
Distilled water to 10.0 ml
INTRODUCTION
Bacterial cells contain endonucleases (enzymes that can cut DNA in nonterminal positions) of high specificity. The purpose of these enzymes within the cell is defensive: to destroy invaders. These enzymes can recognize special nucleotide sequences called restriction sites and confine their activity only to these sites. Most of them can selectively cut at symmetrical base sequences. Under normal conditions, the cell's own DNA is protected from them by modification, accomplished by methylation. Specific bacterial genes are responsible for the production of restriction and modification enzymes.
Restriction enzymes can be used to produce certain DNA fragments with known ends (and lengths). Restriction endonucleases are named according to a specific system of nomenclature. The letters refer to the organism from which the enzyme was isolated. The first letter of the name stands for the genus of the organism. The next two letters represent the second word of the species name. The fourth letter (if there is one) represents the strain of the organism. Roman numerals indicate whether the particular enzyme was the first isolated, the second, or so on. Example: EcoRI--E = Escherichia; co = specific epithet coli; R = strain RY13; I = the first endonuclease isolated.
EcoRI restriction endonuclease can generate six different fragments of linear DNA of lambda phage because it has five recognition sites. The same enzyme recognizes only one cleavage point in simian virus SV40. HindII may cut SV40 at five positions, and lambda at 34. These fragments can then be separated by electrophoresis. Using restriction endonucleases, physical maps of the DNA can be constructed.
The lambda DNA molecule has an unusual structure (Fig.1). Single-stranded segments of DNA containing 12 nucleotides are found at both ends of the phage DNA molecule. The base sequence of these terminal regions, known as cohesive or sticky ends, are complementary to each other. By forming base pairs between the cohesive ends, the linear DNA molecule will circularize. When lambda DNA is incubated at 37oC, the cohesive ends anneal to each other by base-pairing and the DNA molecule is converted to a circular form. When the circular DNA is heated to 65oC, the cohesive ends come apart and the DNA molecular is converted to a linear form. The restoration of the double-helix is called renaturation.
Figure 1. Diagram of the linear and circular forms of the lambda DNA molecule.
The DNA of lambda phage may be divided into three regions (Figs. 1 & 2). The left-hand region includes all the genes whose products are necessary to produce phage head and tail proteins and to package the DNA into the virus. The central regions contain elements involved in integration of the DNA into the E. coli chromosome. The remaining portion of the genome includes the major control regions, the genes necessary for replication and those for cell lysis. Five EcoRI restriction sites occur in the lambda DNA. Cleavage at these sites by EcoRI produces six DNA fragments. Two of these fragments (the fragments derived from the termini) contain cohesive ends and will anneal together under the proper conditions.
Figure 2. Genetic and EcoRI restriction map for lambda DNA.
A first step in the analysis of a DNA molecule frequently involves the determination of its length in nucleotide pairs. Electrophoresis in agarose gels has proven to be a useful tool for this purpose, and for the separation of DNA fragments of different sizes. The length of a given DNA fragment can be determined by comparing its electrophoretic mobility on agarose gels with DNA makers of known lengths. The smaller a DNA fragment, the more rapidly it moves during electrophoresis. As shown in Figure 3, a linear relationship is obtained if the logarithms of the size (in base-pair units) of the DNA fragments are plotted against their electrophoretic mobilities. The length of an unknown DNA fragment is then estimated from this calibration curve. In practice DNA standards and unknown DNAs are electrophoresed on adjacent lanes of the same agarose gel. After electrophoresis, the positions of the standard and unknown DNA bands in the gel are determined and the size of the unknown calculated. The length of DNA is frequently given in base-pairs (bp) for small fragments and kilobase pairs (kb) for larger ones. One kilobase-pair equal 1000 base-pairs.
OBJECTIVES
To introduce the genetic analysis of DNA using restriction enzymes and gel electrophoresis. To determine the lengths of DNA fragments.
[pic]
Figure 3. The distance migrated by six restriction fragments are plotted against the logarithms of their lengths. The length of any unknown can be determined by extrapolation from the standard graph.
Laboratory Session 1
DIGESTION WITH RESTRICTION ENZYME
Digesting Lambda DNA with EcoRI, Hind III, and Pst I
1. Label four 1.5 ml microtubes as follows: C = Control; E = EcoRI; P = Pst I; H = Hind III.
2. Using a V-10 micropipettor (0.5-10 µl), add 4 µl of lambda DNA, 5 µl of restriction buffer, and 1 µl of restriction enzyme to each tube according to the following table. Add only one kind of restriction enzyme to a tube. First add DNA, then restriction buffer, and then enzyme to tubes. Use a fresh pipette tip for restriction buffer and each enzyme. Note: Restriction enzymes are very temperature sensitive. Keep them on ice at all times.
| |Lambda |Restriction | | | |
|Tube |DNA |Buffer |EcoRI |Pst I |Hind III |
| C |4 µl |5 µl |---- |---- |---- |
| E |4 µl |5 µl |1 µl |---- |---- |
| P |4 µl |5 µl |---- |1 µl |---- |
| H |4 µl |5 µl |---- |---- |µl |
2. Close the tube tops and place the tubes into a microcentrifuge, being sure to space them evenly around the inside. Pulse spin the tubes for a few seconds.
3. Place the tubes in a 37oC water bath and incubate for 1 hour.
4. Following incubation, freeze the samples until the next laboratory period.
Pour an agarose gel, wrap the gel in plastic wrap, and store the gel in the refrigerator for use in Laboratory Session 2.
Laboratory Session 2
ELECTROPHORESIS PROCEDURE
Pouring the Agarose Gel
1. Place the Mini-Gel Caster on a level surface. Disengage and slide the movable wall to the open of the caster by turning and lifting the cam peg upward.
2. Place the open end of the gel tray against the fixed wall of the gel caster. Slide the movable wall against the edge of the gel tray and engage the cam peg by turning and pressing downward simultaneously. When the cam peg has dropped into place, turn the peg until resistance is felt. This will seal the edges of the tray for gel casting.
3. Insert the comb into the slot at the end of the gel tray.
4. Make a 0.8% agarose gel solution. Place 30 ml of electrophoretic buffer (1x TBE or TAE) into a 100 ml Erlenmeyer flask or a 100 ml bottle and add 0.24 g of agarose. Gently swirl the flask until the agarose forms a suspension.
5. Heat the agarose in the microwave oven, using 50% power for 30-45 seconds. Be sure to cover flask with plastic wrap or to use a bottle with a loosened cap (to release pressure during heating). Stir frequently during microwaving and stop if the solution starts to boil. The agarose solution should be absolutely clear with no un-melted agarose visible. The correct protocol for microwaving the agarose is to mix the agarose in the buffer, microwave, stop, mix, microwave, etc.
6. Allow the agarose solution to cool to 50oC (check with thermometer). Hot agarose may cause the gel tray and comb to warp or crack.
7. Pour the agarose onto the gel tray until the comb teeth are submerged in gel solution (~20 ml). Remove any air bubbles from the agarose.
8. Allow the gel to cool for 15 minutes. The gel must be completely solidified before the comb is removed or the wells may be improperly formed and the DNA bands distorted.
9. Carefully remove the comb from the solidified gel by lifting it straight out of the gel slowly.
10. Disengage the cam peg by turning and lifting upward. When the gel is solidifying, a light seal is formed between the gasket and the gel. Before moving the wall of the gel caster, carefully lift the gel tray on one side to release the seal or use a spatula to break the seal between the agarose and gasket. Gels can be stored for one week before use. For gel storage, the comb is left in place and the tray containing the gel and comb is wrapped in plastic wrap or placed in a zip lock bag and stored in the refrigerator.
11. Place the gel tray onto the base of the electrophoretic cell so that the sample wells are near the cathode (negative pole or black). The DNA molecules migrate toward the anode (positive pole or red) during electrophoresis.
12. Pour ~275 ml of 1x electrophoretic buffer (TBE or TAE) into the buffer tray. The gel should be completely submerged with the buffer is 2 to 6 mm above the gel surface.
Loading the Samples
1. Thaw the DNA samples. Set the V-10 digital micropipettor to 2.0 µl and add 2 µl of bromophenol blue to each of tube labeled C, E, P, and H. Close tube tops and pulse spin tubes in the microcentrifuge for a few seconds to thoroughly mix the DNA samples and the loading dye. Use a fresh tip with each sample to avoid contamination.
2. Carefully load 10 µl of each sample into separate wells. Be careful not to punch the tip of the pipette through the bottom of the gel.
3. Use a fresh pipette tip for each sample.
|Sample Well No. |Sample |
|1 |DNA from Tube C |
|2 |DNA from Tube E |
|3 |DNA from Tube P |
|4 |DNA from Tube H |
4. Place the lid on the electrophoretic cell carefully so as not to disturb the samples. The lid attaches to the cell in only one orientation. To attach the lid correctly, match the red and black banana jacks on the lid with the red and black banana plugs of the cell.
Electrophoresis
1. Connect the electrophoresis cell to the power supply by inserting the molded two-prong plugs into the power supply’s high voltage output jacks.
2. Press the Power Switch that is located on the side of the power supply. Press the side labeled “I” on the switch.
3. Press the Constant (Const) key to select constant voltage.
4. Use the Scroll Key to enter 100 volts.
5. Press the Run Key to start the run.
6. Electrophorese the samples for ~45 minutes.
7. Stop the run by pressing the Run Key.
8. Press the Power Switch to turn the power supply off, disconnect the power cords from the cell, remove the top of the electrophoresis cell, and remove the gel tray containing the gel.
9. Pour the electrophoretic buffer back into its original container and store it in the refrigerator. Rinse the electrophoresis cell and lid with water.
Staining and Destaining
1. Gloves should be worn to avoid contact with the staining solution and during handling of the gel. Slide the gel from the casting tray onto a piece of plastic wrap.
A. Moisten the gel with a few drops of electrophoretic buffer.
B. Obtain an InstaStainTM ethidium bromide card from the laboratory instructor.
C. Remove the plastic film that covers the unprinted side of the InstaStainTM ethidium bromide card. The unprinted side of the card contains the ethidium bromide that will be used to stain the DNA.
D. Place the unprinted side of the InstaStainTM card on the gel and firmly run your fingers over the entire surface of the card several times.
E. Place the gel casting tray on top of the gel and put a small 50 ml beaker on the gel tray to hold the InstaStainTM card in contact with the gel.
F. After 10 minutes, remove the InstaStainTM card and place the card in the hazardous waste container. View the gel on a UV transilluminator.
G. Make sure you wear UV-blocking goggles, gloves, and a lab coat or long sleeved shirt to protect yourself from the intense ultraviolet light emitted by the transilluminator. This will prevent a severe "sunburn" that can result from even short exposures to the UV light from the transilluminator.
1. Alternative staining procedure with InstaStainTM Methylene Blue.
A. Moisten the gel with a few drops of electrophoretic buffer.
B. Obtain an InstaStainTM Methylene Blue card from the laboratory instructor.
C. Place the unprinted side of the InstaStainTM card on the gel and firmly run your fingers over the entire surface of the card several times.
D. Place the gel casting tray on top of the gel and put a small 50 ml beaker on the gel tray to hold the InstaStainTM card in contact with the gel.
E. After 15 minutes, remove the InstaStainTM card and place the card in the hazardous waste container. Transfer the gel to a staining tray and destain by adding distilled water to slightly submerge the gel. Change the water every 10 minutes until DNA bands become visible.
F. After destaining, transfer the gel to a light box to view bands. You must work carefully because the methylene blue will fade quickly when exposed to light.
3. A photograph of the gel will be taken using the photodocumentation system.
DETERMINING THE LENGTH OF A DNA MOLECULE
1. Measure the migration distance in millimeters for each Hind III band from the predigested DNA. Measure from the front edge of the sample well to the front edge of the band. Enter the distances in Table 1.
2. Obtain a piece of semilog graph paper. Mark the x-axis in millimeters. This axis represents the migration distance. Label the axis.
3. Fragment size (in base pairs) is graphed along the y-axis. Assume that the first section or cycle of semilog pager represents 0-1000 base pairs, the second represents 1000-10,000 base pairs, and the third cycle represents 10,000-100,000 base pairs (recall logarithms differ by the power of 10).
4. Plot the migration distance for each band of the Hind III predigest DNA against the fragment sizes given in Table 1. Connect the data points with a line. This will serve as a standard curve.
5. Measure the migration distance in millimeters for each EcoRI band and enter in Table 1. Measure the migration distance for each Pst I band and enter in Table 1.
6. Using the standard curve, determine the sizes of the fragments of phage lambda DNA digested with EcoRI and Pst I. This is accomplished by locating on the x axis the distance migrated by the fragment. Using a ruler, draw a vertical line from this point to its intersection with the data line. Now extend a horizontal line from this point to the y axis. This gives the size of the fragment. Note: This technique is not exact--you should expect as much as 10% to 15% error.
7. Enter the results in estimated base-pair column in Table 1.
8.
Table 1. Base Pair Sizes of Lambda DNA Fragments Generated by Restriction Digest
[pic]
9. For which fragment sizes was your graph most accurate? For which fragment sizes was your graph least accurate? What does this tell you about the resolving ability of agarose gel electrophoresis?
ESTIMATION OF SIZE OF RESTRICTION FRAGMENTS BY REGRESSION ANALYSIS – GRAPHING IN MICROSOFT EXCEL (See Appendix C)
1. Open Microsoft Excel by clicking on the Excel icon.
2. Enter your data from your Hind III predigest DNA in Columns A (distance migrated) and Column B (fragment size).
3. After entering your data, click the Charting Icon on the toolbar, or select Chart in the Insert menu. The Chart wizard will open.
4. In the chart window select XY (Scatter) chart type. Continue by clicking Next.
5. Select the columns that you want to plot. Continue by clicking Next.
6. Select the Titles tab and fill in the information for the x-axis title as Distance (mm). Fill in the y-axis as Base Pairs (bp). Select the Gridlines tab and place checks for major and minor gridlines for both x- and y-axes. Select the Legend tab and remove the check by Show Legend. Continue by clicking Finish.
7. The graph should have been inserted into the spreadsheet. Place your cursor on the y-axis and double click. Select Scale tab and under the Value (Y) Axis Scale enter the size of the smallest fragment as Minimum and the size of the largest fragment as Maximum. Place a check next to Logarithmic scale. Hit OK.
8. Place your cursor of the x-axis and double click. Select Scale tab and under the Value (X) Axis Scale enter the distance the smallest fragment migrated as Minimum and the distance the largest fragment migrated as Maximum. Hit OK.
9. Next, add a trendline. Under Chart in the toolbar, select Add Trendline. In the trendline window, select exponential. Under Options, select R2 value and show equation. Click OK. Your graph should now show a linear line, regression equation, and R2 value. The regression equation shows the equation for the line generated by the data
10. You now have both a graphical representation of the relationship between the size and the migration distance of your DNA fragments, and the mathematical description of that relationship. Since you can measure X (migration distance), you can now calculate accurately the value of Y (size). Use the regression equation to estimate the size of your restriction fragments and enter the information in Table 2.
11. See Appendix C for a discussion of regression analysis.
Table 2. Base Pair Sizes of Lambda DNA Fragments Estimated by Regression Analysis
| |Lambda DNA |Lambda DNA |Lambda DNA |Lambda DNA |
| |No Enzyme – |EcoRI – |Pst I – |Hind III – |
|Band |Est. bp |Est. bp |Est. bp |Est. bp |
|1 | | | | |
|2 | | | | |
|3 | | | | |
|4 | | | | |
|5 | | | | |
|6 | | | | |
SUGGESTED READINGS
Cohen, S. N. 1975. The manipulation of genes. Sci. Amer. 233:24-33.
Grobstein, C. 1977. The recombinant DNA debate. Sci. Amer. 237:22-33.
Helling, R. B., H. M. Goodman, and H. W. Boyer. 1974. Analysis of EcoRI fragments of DNA from lamboid bacteriophage and other viruses by a agarose gel electrophoresis. J. Virol. 14:1235.
White, R. and J. -M. Lalouel. 1988. Chromosome mapping with DNA markers. Sci. Amer. 258:40-48.
EXERCISE 9
DNA FINGERPRINTING (PROFILING)
Laboratory Session One
Items for each Student Workstation
EcoRI/PstI enzyme mixture - 1 tube (80 µl)
Sterile Pipet tips
Micropipet
Color coded microtubes: green, blue, orange, violet, red, yellow - 1 each
Lab marker
Waste container
Styrofoam microtube rack
Ice bucket with ice
Gel trays
Sample well-forming combs (1/group)
1x TAE electrophoresis buffer
Items for Instructor’s Workstation
Crime Scene DNA with buffer, rehydrated - 1 vial
Suspect 1 DNA with buffer, rehydrated - 1 vial
Suspect 2 DNA with buffer, rehydrated - 1 vial
Suspect 3 DNA with buffer, rehydrated - 1 vial
Suspect 4 DNA with buffer, rehydrated - 1 vial
Suspect 5 DNA with buffer, rehydrated - 1 vial
Water bath - (37 °C) 1/class
Balances
Agarose
Microcentrifuges
Laboratory Session Two
Items for each Student Workstation
Agarose gel
Digested DNA samples from crime scene and suspects
DNA sample loading dye
Marking pen
Pipet tips
Micropipets
Waste container
Styrofoam microtube rack
Gel staining tray
BioRad Electrophoretic Unit
Power Supply
1x TAE Electrophoresis buffer 275 ml gel/box
Items for Instructor’s Workstation
InstaStainTM ethidium bromide card or InstaStainTM methylene blue card or Bio-Safe DNA stain - 1x solution 500 ml
HindIII DNA markers
Laboratory Session Three
Photodocumentation system
Millimeter ruler
Semi-log graph paper
INTRODUCTION
No two individuals, except identical twins, have exactly the same DNA sequence. This has lead to the development of DNA fingerprinting (also known as DNA profiling or DNA typing) that has been employed to identify a criminal using DNA samples from a crime scene, to exonerate innocent persons, to determining paternity, and to screen for genetic diseases. DNA obtained from body tissues, body fluids (blood, semen, saliva, or urine), or hair is digested with restriction endonucleases and then separated into different-sized fragments using electrophoresis. This profile of DNA fragments looks like a “bar code” and yields a distinctive and unique "DNA fingerprint" for each the individual.
Slight but unique differences in the banding pattern of restriction fragments from different individuals are observed when the DNA is digested with restriction endonucleases and separated by electrophoresis. These variations in the DNA are called restriction fragment length polymorphisms (RFLPs; riff-lips in biotech jargon) and occur because individuals possess variable restriction sites so that two pieces of DNA from separate individuals may have different fragment lengths when their DNA is cut by the same restriction enzyme. Comparing the different-sized DNA fragments of two samples provides very strong evidence about whether or not the two samples came from a single individual.
In this laboratory exercise, you will explore how DNA samples can be distinguished from each other based on variations in the nucleotide sequence of the DNA. This technology is commonly used in forensic science and is now employed in numerous television shows each week to solve baffling crimes. A heinous crime has just been committed and you and your team of experts have been assigned the task of determining “who done it.” You have collected samples of DNA from a crime scene and 5 suspects that must be analyzed to determine which if any of the suspects may be guilty.
Laboratory Session 1
1. Label reaction tubes.
A. Obtain one each of the following colored microtubes. Label the 5 colored microtubes as follows:
Green CS (crime scene)
Blue S1 (suspect 1)
Orange S2 (suspect 2)
Violet S3 (suspect 3)
Red S4 (suspect 4)
Yellow S5 (suspect 5)
B. Label the tubes with your name and lab period number. The restriction digests will occur in these tubes. These tubes may now be kept in your rack.
[pic]
2. Pipet 10 µl of each DNA sample from the stock tubes located at the Instructor’s Workstation and transfer to the corresponding colored microtubes. Use a separate tip for each DNA sample. Make sure the sample is transferred to the bottom of the tubes.
[pic]
3. Using a new tip for each DNA sample, add 10 (L of “enzyme” containing the EcoRI and PstI restriction enzymes to the crime scene and 5 suspect DNA samples (each of these tubes should already contain 10 (L DNA).
4. Cap the tubes and mix the components by gently flicking the tubes with your finger. Pulse spin the samples in the microcentrifuge to collect all the liquid in the bottom of the tube. (Be sure the tubes are in a BALANCED arrangement in the rotor).
5. Place the tubes in the floating rack and incubate them in the water bath at 37 °C for 1 hour. After the incubation, store the DNA digests in the refrigerator until the next lab period.
Pouring the Agarose Gel
1. While the restriction digest is taking place in the water bath, pour an agarose gel that will be used during the next laboratory period.
2. Place the Mini-Gel Caster on a level surface. Disengage and slide the movable wall to the open of the caster by turning and lifting the cam peg upward.
3. Place the open end of the gel tray against the fixed wall of the gel caster. Slide the movable wall against the edge of the gel tray and engage the cam peg by turning and pressing downward simultaneously. When the cam peg has dropped into place, turn the peg until resistance is felt. This will seal the edges of the tray for gel casting.
4. Insert the comb into the slot at the end of the gel tray.
5. Make a 1.0% agarose gel solution. Place 30 ml of 1x TAE electrophoretic buffer (1x into a 100 ml bottle and add 0.3 g of agarose. Gently swirl the flask until the agarose forms a suspension.
6. Heat the agarose in the microwave oven, using 50% power for 30-45 seconds. Be sure to cover flask with plastic wrap or to use a bottle with a loosened cap (to release pressure during heating). Stir frequently during microwaving and stop if the solution starts to boil. The agarose solution should be absolutely clear with no un-melted agarose visible. The correct protocol for microwaving the agarose is to mix the agarose in the buffer, microwave, stop, mix, microwave, etc.
7. Allow the agarose solution to cool to 50oC (check with thermometer). Hot agarose may cause the gel tray and comb to warp or crack.
8. Pour the agarose onto the gel tray until the comb teeth are submerged in gel solution (~20 ml). Remove any air bubbles from the agarose.
9. Allow the gel to cool for 15 minutes. Disengage the cam peg by turning and lifting upward. When the gel is solidifying, a light seal is formed between the gasket and the gel. Before moving the wall of the gel caster, carefully lift the gel tray on one side to release the seal or use a spatula to break the seal between the agarose and gasket. Gels can be stored for one week before use. For gel storage, the comb is left in place and the tray containing the gel and comb is wrapped in plastic wrap or placed in a zip lock bag and stored in the refrigerator.
Laboratory Session 2
Electrophoresis of DNA Samples
1. Obtain your agarose gel from the refrigerator. Remove the plastic wrap and carefully remove the comb from the solidified gel by lifting it straight out of the gel slowly.
2. Place the agarose gel in the electrophoresis apparatus. Pour 1x TAE buffer in the gel box until it just covers the wells. This will require approximately 275 ml of buffer. Check that the wells of the agarose gels are near the black (-) electrode and the base of the gel is near the red (+) electrode.
3. Remove your digested DNA samples from the refrigerator. Pulse spin the tubes in the microcentrifuge to bring all of the liquid into the bottom of the tube.
4. Using a separate tip for each sample, add 5 µl of loading dye "LD" into each tube. Close the caps on the tubes and mix by gently flicking the tube with your finger. Pulse spin the tubes in the microcentrifuge to bring the contents to the bottom
5. Using a separate pipet tip for each sample, load your gel as follows:
Lane 1: HindIII DNA size marker, clear tube 10 µl
Lane 2: CS, green tube 20 µl
Lane 3: S1, blue tube 20 µl
Lane 4: S2, orange tube 20 µl
Lane 5: S3, violet tube 20 µl
Lane 6: S4, red tube 20 µl
Lane 7: S5, yellow tube 20 µl
6. Secure the lid on the gel box. The lid will attach to the base in only one orientation: red-to-red and black-to-black. Connect electrical leads to the power supply.
7. Turn on the power supply. Set it for 100 V and electrophorese the samples for 60 minutes.
8. When the electrophoresis is complete, turn off the power and remove the lid from the gel box. Using gloves, carefully remove the gel tray and the gel from the gel box. Be careful, the gel is very slippery! Nudge the gel off the gel tray with your thumb and carefully slide it onto a piece of plastic wrap and stain with ethidium bromide for 10 minutes using the following procedure:
A. Moisten the gel with a few drops of electrophoretic buffer.
B. Obtain an InstaStainTM ethidium bromide card from the laboratory instructor.
C. Remove the plastic film that covers the unprinted side of the InstaStainTM ethidium bromide card. The unprinted side of the card contains the ethidium bromide that will be used to stain the DNA.
D. Place the unprinted side of the InstaStainTM card on the gel and firmly run your fingers over the entire surface of the card several times.
E. Place the gel casting tray on top of the gel and put a small 50 ml beaker on the gel tray to hold the InstaStainTM card in contact with the gel.
F. After 10 minutes, remove the InstaStainTM card and place the card in the hazardous waste container. View the gel on a UV transilluminator.
9. Alternative staining procedure with InstaStainTM Methylene Blue.
A. Moisten the gel with a few drops of electrophoretic buffer.
B. Obtain an InstaStainTM Methylene Blue card from the laboratory instructor.
C. Place the unprinted side of the InstaStainTM card on the gel and firmly run your fingers over the entire surface of the card several times.
D. Place the gel casting tray on top of the gel and put a small 50 ml beaker on the gel tray to hold the InstaStainTM card in contact with the gel.
G. After 15 minutes, remove the InstaStainTM card and place the card in the hazardous waste container. Transfer the gel to a staining tray and destain by adding distilled water to slightly submerge the gel. Change the water every 10 minutes until DNA bands become visible.
H. After destaining, transfer the gel to a light box to view bands. You must work carefully because the methylene blue will fade quickly when exposed to light.
10. Alternative staining procedure with Bio-Safe DNA stain.
A. Add 60 ml of Bio-Safe DNA stain to the tray. Let the gel stain overnight, with shaking for best results.
B. Pour off the Bio-Safe DNA stain into a bottle or another appropriate container and destain the gel with 60 ml of water for ~15 minutes.
C. Pour the water out of the staining tray. Ask the instructor how to properly dispose of the stain.
11. Pour the electrophoretic buffer back into its original container and store it in the refrigerator. Rinse the electrophoresis cell and lid with water.
Laboratory Session 3
DETERMINING THE LENGTH OF A DNA MOLECULE
1. Measure the migration distance in millimeters for each Hind III band from the predigested DNA. Measure from the front edge of the sample well to the front edge of the band. Enter the distances in Table 1.
2. Obtain a piece of semilog graph paper. Mark the x-axis in millimeters. This axis represents the migration distance. Label the axis.
3. Fragment size (in base pairs) is graphed along the y-axis. Assume that the first section or cycle of semilog pager represents 0-1000 base pairs, the second represents 1000-10,000 base pairs, and the third cycle represents 10,000-100,000 base pairs (recall logarithms differ by the power of 10).
4. Plot the migration distance for each band of the Hind III predigest DNA against the fragment sizes given in Table 1. Connect the data points with a line. This standard curve will be used to estimate the sizes of the crime scene and suspect restriction fragments.
5. Measure the migration distance in millimeters for each all crime scene and suspect restriction fragments and enter the data in Table 1.
6. To estimate the size of an unknown crime scene or suspect fragment, find the distance that fragment traveled. Locate that distance on the x-axis of your standard graph. From that position on the x-axis, read up to the standard line, and then follow the graph line to over to the y-axis. You might want to draw a light pencil mark from the x-axis up to the standard curve and over to the y-axis showing what you’ve done. Where the graph line meets the y-axis, this is the approximate size of your unknown DNA fragment. Do this for all crime scene and suspect fragments. Enter the results in estimated base-pair column in Table 1
7. Compare the fragment sizes of the suspects and the crime scene. Is there a suspect that matches the crime scene?
Table 1. Restriction Fragments from crime scene and suspect DNA
| | | | | | | | |
| |Lambda/Hind III |Crime Scene |Suspect 1 |Suspect 2 |Suspect 3 |Suspect 4 |Suspect 5 |
| |Size marker | | | | | | |
| | | |
|Band |Distance |Actual |
| |(mm) |Size (bp) |
|LB/Amp: +DNA | | |
|LB/Amp/Ara: +DNA | | |
|LB/Amp: -DNA | | |
|LB: -DNA | | |
1. Transformation Efficiency. Transformation efficiency is expressed as the number of antibiotic-resistant or fluorescent colonies per microgram (µg) of pGLO.
2. Determine the total amount of DNA:
The total amount of DNA we began with is equal to the product of the concentration and the total volume used or
DNA (µg) = (Concentration of DNA in µg/µl) x Volume of DNA in µl)
You used 10 µl of pGLO at a concentration of 0.03 µg/µl. Calculate the total amount of DNA used in this experiment: _____ µg
This number will be multiplied by the fraction of DNA used to determine the total amount of DNA spread on the agar plate.
3. Determine the fraction of DNA spread onto the agar plates:
a. Since not all the DNA you added to the bacterial cells will be transferred to the agar plate, you need to find out what fraction of the DNA was actually spread on the agar plate. To do this, divide the volume of DNA you spread n the plate by the total volume of liquid in the test tube containing the DNA. A formula for this statement is:
Fraction of DNA used = Volume spread on agar plate
Total volume in tube
b. You spread 200 µl of cells containing DNA from a tube containing a total volume of 510 µl (250 µl CaCl2 + 10 µl pGLO + 250 µl of Luria broth
______ Fraction of DNA
c. This number will be multiplied by the amount of DNA used to calculate the amount of DNA spread on an agar plate.
d. Micrograms of DNA spread on agar plate:
pGLO DNA spread (µg) = Total mount of DNA used (µg) x fraction of DNA
_______ µg
Transformation efficiency = Total number of cells growing on the agar plate
Amount of DNA spread on the agar plate
| | |µg DNA Spread |Transformation |
|Agar Plate |# of Colonies |on Agar Plate |Efficiency |
|LB/Amp: +DNA | | | |
|LB/Amp/Ara: +DNA | | | |
SUGGESTED READINGS
Hanahan, D. 1983. Studies on transformation of Escherichia coli with plasmids. J. Mol Biol. 166:557.
Hotchkiss, R. D. and M. Gabor. 1970. Bacterial transformation, with special reference to recombination process. Ann. Rev. Genet. 4:193-224.
Novick, R. P. 1980. Plasmids. Sci. Amer. 243:102-127.
Smith, H. O., D. B. Danner, and R. A. Deich. 1982. Genetic transformation. Ann. Rev. Biochem. 50:41-68.
EXERCISE 11
BIOINFORMATICS
MATERIALS
Internet capability
INTRODUCTION
Bioinformatics is a newly developing field that that applies computer and information science to solve biological problems. The need for such capabilities has been precipitated by the exponential explosion of publicly available biological data from various genome projects, such as the Human Genome Project. The rapid expansion of biological information has necessitated the development of new ways of storing and analyzing genetic information and databases. These databases are rapidly growing collections of DNA and protein sequences that are submitted by researchers worldwide.
Bioinformatics organizes data in a way that allows researchers to access existing information and to submit new data as they are produced. Bioinformatics also seeks to develop tools and resources that aid in the analysis of data and the interpretation of results in a biologically meaningful manner. These tools are based on algorithms, which are problem-solving methods performed by computers. Algorithms are used in numerous ways, from generating amino acid sequences from DNA sequence to predicting the complexities of protein structure. Traditionally, biological studies examined individual systems in detail, and frequently compared them with a few that are related. In bioinformatics, scientists can conduct global analyses of all the available data
The National Center for Biotechnology Information (NCBI) is a federally funded repository for a large amount of the genetic data and also provides tools for use in analyzing the data. This laboratory exercise is designed to acquaint you with a few of these different tools and how they are utilized to manage genomic data and provide information on the structure and function of a gene or gene product. One of the most popular tools for comparing a newly derived DNA or protein sequence with the information in the databases is BLAST, an acronym for Basic Local Alignment Search Tool. BLAST is a set of similarity search programs designed to explore all of the available sequence databases. It will match several small subsequences within your search sequence to the sequences within your search sequence to the sequences in the database. The BLAST output describes sequences that have regions of similarity to your sequence, and provides a variety of links to those sequences and other analysis programs.
Protein Sequence Alignment
In this exercise you will search for protein (amino acid) sequences in the databases that are similar to the query sequence. The database is searched for proteins similar to your query and the computer aligns your protein (query) sequence with that of all other proteins in the database and determines the ones with the best fit. The process of aligning two or more sequences such that regions of similarity between molecules are most apparent is called sequence alignment. By comparing the protein to known sequences in the databases, you may be able to develop a testable hypothesis regarding the structure and function of the protein.
Your instructor will assign each group one of the following amino acid sequences. The amino acids are indicated with their single-letter symbols (Figure 1).
A. GPRNCIGQKFATLEEKTVLS
B. GPRNCIGQTFAMSEMKVALA
C. GIRVCLGEVLAKMELFLFLA
D. GPRSCVGEMLARQELFFFTA
E. GKRRCLGEVIGRWEVFLFLA
F. GKRKCIGETIGRLEVFLFLA
|One-Letter Symbol|Three-Letter | |
| |Abbreviation |Full Name |
|A |Ala |Alanine |
|R |Arg |Arginine |
|N |Asn |Asparagine |
|D |Asp |Aspartic acid |
|C |Cys |Cysteine |
|Q |Gln |Glutamine |
|E |Glu |Glutamic acid |
|G |Gly |Glycine |
|H |His |Histidine |
|I |Ile |Isoleucine |
|L |Leu |Leucine |
|K |Lys |Lysine |
|M |Met |Methionine |
|F |Phe |Phenylalanine |
|P |Pro |Proline |
|S |Ser |Serine |
|T |Thr |Threonine |
|W |Trp |Tryptophan |
|Y |Try |Tyrosine |
|V |Val |Valine |
|B |Asx |Aspartic acid |
|Z |Glx |Glutamine |
Open . This web site is linked to a large number of DNA and protein sequence databases. For example, you can use PubMed to find journal articles on any topic remotely related to medicine, Entrez to find DNA or protein sequences by name or OMIM (Online Mendelian Inheritance in Man) to find information about specific human genetic disorders.
1. On the navigation bar, select BLAST (Basic local Alignment Search Tool). This is an algorithm to align sequences and to select the ones that are closest to the on you will be entering for comparison.
2. Under “Protein Blast” select Standard protein-protein BLAST (blastp).
3. Type your assigned amino acid sequence in the search field.
4. In the database pull menu, choose nr (non-redundant). This means that you will be scanning through the whole database, skipping any duplicates that are present in the database. The “set subsequence” fields can be left open, and the “Do CD-Search” box is unchecked.
5. Click on the BLAST icon to start your search. This will take you to another page that will display your Request ID number and a message giving you the approximate amount of time it will take before the search is completed.
6. Click on the FORMAT button, which will open a new window in your browser. If you results are not ready, the page will be largely empty, with a message that it will automatically reload shortly.
7. Print out the results.
8. Answers the questions at the end of the laboratory exercise.
Interpreting your blastp search
The BLAST output shows you what proteins sequences share regions of identity with your protein query, what organisms the matching sequences come from, and how closely they match. The information is given to you in three different ways: a graphical, color-coded view; a set of one-line summaries; and a detailed match description that includes alignments and comparisons of your query and matching sequence. These results allow you to assign a function to your new protein.
The color-coded view is at the top of the page, and consists of a series of different colored lines. The red bar at the top of the figure represents your query sequence. The different colored bars below that line represent the matches that were found in the databases. Very close matches are shown in red, relatively close matches in lavender, and so on, with black regions indicating possible low matches and white regions indicating areas of no sequence identity.
Below this figure, a slightly more detailed version of the same information is presented in a series of one-line summaries. Scroll down the page to this region of the output. Each line has three parts: a hyperlink to the GenBank entry where the matching sequence can be found; the name of the organism that the sequence came from; and a set of values indicating how closely the sequence matches with the query. A high score (bit) value and/or a low E-value (expect value) indicate close matches. A bit score is a measure of similarity between the hit and the query. The E-value is a statistical measure of the likelihood of similar sequences occurring randomly. Biologically significant hits will tend to have E-values much less than 1.0. The larger the E-value the greater the chance that the similarity between the hit and the query is due to mere coincidence.
Scroll down to the third section of the report. The amino acid alignment is presented as letters representing the amino acid repeated between the two sequences. In some position there is a “+” sign between the two sequences. This represents a site where the amino acids are different but very similar; for example an acidic amino acid in the query sequence lines up with a different, but still acidic amino acid in the subject sequence. The summary at the top of the alignment show the number of amino acids that are identical and the number that is similar.
DNA Sequence Similarity
Your instructor will assign each group one of the following nucleic acid sequences.
A. GCCAGAGATGAATGTGGTTTTGAAAATGTCAAACCAGACCAATGGGTGA
B. TCCAGTGTTCACAGTAAGATGTACTCAGGCCAGTCCATGGGCGGCCGTGG
C. GAAAAAGAAATATTTGAAAGCTGTGTCTGTAAATTGATGGCTAACAAAAC
D. CCCGTGAAAAAGAGCCGCCTGCAGCTGCTGGGGGCCACTTGCATGTTCGT
Go back to the NCBI homepage.
1. Choose BLAST on the Navigation Bar
2. Under “Nucleotide BLAST,” click on “Standard nucleotide-nucleotide BLAST (blastn).”
3. Type your assigned nucleic acid sequence in the search field.
4. Leave “set subsequences” open and click on BLAST.
5. Click on FORMAT.
6. Print out the results.
7. Answer the questions at the end of the laboratory exercise.
Interpreting your blastn search
The format is similar to that for blastp, and the information is given to you in three different ways: a graphical, color-coded view; a set of one-line summaries; and a detailed match description that includes alignments and comparison of your query and the matching sequence.
The color-coded view is at the top of the page, and consists of a series of different colored lines. The red bar at the top of the figure represents your query sequence. The different colored bars below that line represent the matches that were found in the databases. Very close matches are shown in red, relatively close matches in lavender, and so on, with black regions indicating possible low matches and white regions indicating areas of no sequence identity.
Below this figure, a slightly more detailed version of the same information is presented in a series of one-line summaries. Scroll down the page to this region of the output. Each line has three parts: a hyperlink to the GenBank entry where the matching sequence can be found; the name of the organism that the sequence came from; and a set of values indicating how closely the sequence matches with the query. A high score value and/or a low E value indicate close matches.
Scroll down to the third section of the report. On this page you can see an alignment of sections of the query and match sequences. The vertical lines represent nucleotides that are identical in each of the sequences. The number of the “Subject,” or match sequences, refers to the numbering of the sequence in its GenBank entry. There is also a listing of percent sequence identity.
QUESTIONS
Protein Sequence Alignment
1. What is the name of the protein and species whose sequence matches the one you were given (You may need to examine a few of the best E-values to determine the protein name).
________________________________________________________________
2. What is the E-value for this match? ____________________________________
3. What is the function of the protein? ____________________________________
DNA Sequence Similarity
1. What is the name of the gene and species whose sequence matches the one you were given (You may need to examine a few of the best E-values to determine the protein name).
________________________________________________________________
2. What is the E-value for this match? ____________________________________
3. What other proteins and species also came up as matches to this sequence?
_________________________________________________________________
_________________________________________________________________
_________________________________________________________________
_________________________________________________________________
4. Why would many different organisms have some similar or identical genes?
_________________________________________________________________
_________________________________________________________________
_________________________________________________________________
5. Which genes came up as poor matches to your input sequence and what are their E-values?
_________________________________________________________________
_________________________________________________________________
_________________________________________________________________
6. Where can you find more information about the sequences that matched the input sequence?
_________________________________________________________________
_________________________________________________________________
_________________________________________________________________
_________________________________________________________________
EXERCISE 12
DNA AMPLIFICATION BY POLYMERASE CHAIN
REACTION (PCR)
EQUIPMENT
Perkin Elmer Thermal Cycler 2400
Microcentrifuge
BioRad Electrophoretic Unit
Power Supply
Gel casting trays (1/group)
Mini-Gel Caster (1/group)
Sample well-forming combs (1/group)
Balances
Adjustable Micropipettors with tips
Photodocumentation unit
SUPPLIES AND MATERIALS
Laboratory Session 1
0.2 ml PCR tubes (1/student)
Lambda DNA Amplification Kit (Carolina Biological Supply Company)
Ice bath (1/lab table)
Waterproof Marking Pens (1/group)
Laboratory Session 2
InstaStainTM ethidium bromide card
Agarose
100 ml Erlenmeyer flask (1/group)
Plastic wrap
Thermometer
PCR molecular weight markers
Staining tray
TAE or TBE Buffer
Loading dye
Laboratory Session 3
Transilluminator
Plastic wrap
50X TAE BUFFER
Tris 242.0 g
Glacial acetic acid 57.1 ml
500 mM EDTA 100.0 ml
Dissolve Tris in 600 ml water. Add acetic acid and EDTA and then add distilled water to make 1 liter
INTRODUCTION
The polymerase chain reaction (PCR) is an extremely powerful procedure that allows the in vitro amplification of a selected DNA sequence several million times. PCR actually mimics the process used by cells to replicate their DNA. During cell division, enzymes called DNA polymerases make a copy of the cellular DNA. DNA polymerase requires template DNA, a supply of the four deoxyribonucleotides, and a primer. During cellular DNA replication, a RNA polymerase (primase) synthesizes a short stretch of RNA complementary to one of the DNA strands at the start site of replication. This primer acts as an attachment site for the DNA polymerase, which then produces the complementary DNA strand. During PCR, short synthetic sequences of single-stranded DNA (20-30 nucleotides) serve as primers. Two different primers are used to bracket the target regions to be amplified. One primer is complementary to one DNA strand at the beginning of the target regions while a second primer is complementary to a sequence on the opposite DNA strand at the end of the target region. Thus, these synthetic oligonucleotides define the sequence to be amplified and sever to prime enzymatic amplification of the intervening nucleotide sequence.
The PCR vial contains all the necessary components for DNA replication: a piece of duplex DNA, large quantities of the four nucleotides, large amounts of the primers, and DNA polymerase. The polymerase was isolated from the bacterium Thermus aquaticus that inhabits hot springs. This enzyme, called the Taq polymerase, is very heat-stable and remains active despite repeated heating during the many cycles of PCR amplification. The PCR cycle is composed of three major steps: In step one, the DNA to be amplified is denatured into single strands. In step two, short single-stranded primers pair with the nucleotides adjacent to the region to be amplified. These primers are made synthetically, and are complementary to the nucleotide sequence flanking the region to be amplified. In step three, DNA polymerase begins at the primers and synthesizes a complementary DNA strand. Repeated cycles amplify the original DNA sequence by more than a million times.
OBJECTIVE
In this exercise, students will utilize PCR to amplify a 1106-base pair sequence from the genome of the lambda bacteriophage. Following amplification, the samples will be electrophoresed in a 1.0% agarose gel.
Laboratory Session 1
Polymerase Chain Reaction
1. Keep the lambda DNA and primer mixture in a beaker of crushed ice during the experiment.
2. Use a permanent marker to label the cap (not side) of the 0.2 ml PCR tube with your initials. This tube contains a Ready-To-Go BeadTM at the bottom. The Ready-To-Go BeadTM contains the Taq polymerase, deoxyribonucleotide triphosphates, MgCl2, and buffer in a freeze-dried pellet
3. Tap the tube on a counter top to isolate the ready to go bead to the bottom of the tube.
4. Using a V-200 micropipettors, add 20 (L of the primer mixture to the labeled PCR tube.
5. Using a V-10 micropipettor, add 10 (L of lambda DNA solution to the tube, close the tube cap tightly and mix the two solutions well by flicking the tube with your finger. Make sure the PCR bead goes into solution. The mixture should appear milky in color at first. Gently mix the contents in the tubes until it turns clear. Tap the tube gently on the countertop to consolidate the contents to the bottom of the tube.
6. Place the PCR tube on ice in the thermal cycler tray and record the tray number for your PCR tube. The DNA thermal cycling should be started as soon as possible to prevent nonspecific priming that could occur if the PCR reactions sit to long.
7. The instructor will place the tubes in the automated thermal cycler that has been programmed with the following step file:
96oC - 1 minute
58oC - 1 minute, 1 cycle
link to:
96oC - 30 seconds
58oC - 1 minute, 30 cycles
link to:
58oC - 10 minutes
8. After completion of the amplification process, store the samples at -20oC until the next laboratory period.
Laboratory Session 2
ELECTROPHORESIS PROCEDURE
Pouring the Agarose Gel
1. Place the Mini-Gel Caster on a level surface. Disengage and slide the movable wall to the open of the caster by turning and lifting the cam peg upward.
2. Place the open end of the gel tray against the fixed wall of the gel caster. Slide the movable wall against the edge of the gel tray and engage the cam peg by turning and pressing downward simultaneously. When the cam peg has dropped into place, turn the peg until resistance is felt. This will seal the edges of the tray for gel casting.
3. Insert the comb into the second slot from the end of the tray.
4. Make a 1.0% agarose gel solution. Place 30 ml of electrophoretic buffer (1x TBE or TAE) into a 100 ml Erlenmeyer flask or a 100 ml bottle and add 0.30 g of agarose. Gently swirl the flask until the agarose forms a suspension.
5. Heat the agarose in the microwave oven, using 50% power for 30-45 seconds. Be sure to cover flask with plastic wrap or to use a bottle with a loosened cap (to release pressure during heating). Stir frequently during microwaving and stop if the solution starts to boil. The agarose solution should be absolutely clear with no un-melted agarose visible. The correct protocol for microwaving the agarose is to mix the agarose in the buffer, microwave, stop, mix, microwave, etc.
6. Allow the agarose solution to cool to 50oC (check with thermometer). Hot agarose may cause the gel tray and comb to warp or crack.
7. Pour the agarose onto the gel tray until the comb teeth are submerged in gel solution (~20 ml). Remove any air bubbles from the agarose.
8. Allow the gel to cool for 15 minutes. The gel must be completely solidified before the comb is removed or the wells may be improperly formed and the DNA bands distorted.
9. Carefully remove the comb from the solidified gel by lifting it straight out of the gel slowly.
10. Disengage the cam peg by turning and lifting upward. When the gel is solidifying, a light seal is formed between the gasket and the gel. Before moving the wall of the gel caster, carefully lift the gel tray on one side to release the seal or use a spatula to break the seal between the agarose and gasket. Gels can be stored for one week before use. For gel storage, the comb is left in place and the tray containing the gel and comb is wrapped in plastic wrap or placed in a zip lock bag and stored in the refrigerator.
11. Place the gel tray onto the base of the electrophoretic cell so that the sample wells are near the cathode (negative pole or black). The DNA molecules migrate toward the anode (positive pole or red) during electrophoresis.
12. Pour ~275 ml of 1x electrophoretic buffer (TBE or TAE) into the buffer tray. The gel should be completely submerged with the buffer 2 to 6 mm above the gel surface.
Loading the Samples
1. Thaw the DNA samples. Using the V-10 micropipettor add 3 µl of loading dye to your PCR reaction tube and mix the content by gently shaking or flicking the tube with your finger,
2. Use a V-200 micropipettor and add 20 µl of PCR/loading dye sample into your assigned well of an agarose gel. Expel any air in the tip before loading and be careful not to punch the tip of the pipette through the bottom of the sample well.
3. Obtain a sample of the size marker DNA from your teaching assistant and load 20 µl in one lane of your gel using the V-200 micropipettor.
4. Place the lid on the electrophoretic cell carefully so as not to disturb the samples. The lid attaches to the cell in only one orientation. To attach the lid correctly, match the red and black banana jacks on the lid with the red and black banana plugs of the cell.
Electrophoresis
1. Connect the electrophoresis cell to the power supply by inserting the molded two-prong plugs into the power supply’s high voltage output jacks.
2. Press the Power Switch that is located on the side of the power supply. Press the side labeled “I” on the switch.
3. Press the Constant (Const) key to select constant voltage.
4. Use the Scroll Key to enter 100 volts.
5. Press the Run Key to start the run.
6. Electrophorese the samples for ~45 minutes. Adequate separation will have occurred when the bromophenol blue dye front has moved at least 50 mm from the wells.
7. Stop the run by pressing the Run Key.
8. Press the Power Switch to turn the power supply off, disconnect the power cords from the cell, remove the top of the electrophoresis cell, and remove the gel tray containing the gel.
9. Pour the electrophoretic buffer back into its original container and store it in the refrigerator. Rinse the electrophoresis cell and lid with water.
Staining and Destaining
1. Using gloves, remove the gel from the casting tray and place it on a piece of plastic wrap and stain with ethidium bromide for 10 minutes using the following procedure:
A. Moisten the gel with a few drops of electrophoretic buffer.
B. Obtain an InstaStainTM ethidium bromide card from the laboratory instructor.
C. Remove the plastic film that covers the unprinted side of the InstaStainTM ethidium bromide card. The unprinted side of the card contains the ethidium bromide that will be used to stain the DNA.
D. Place the unprinted side of the InstaStainTM card on the gel and firmly run your fingers over the entire surface of the card several times.
E. Place the gel-casting tray on top of the gel and put a small 50 ml beaker on the gel tray to hold the InstaStainTM card in contact with the gel.
F. After 10 minutes, remove the InstaStainTM card and place the card in the hazardous waste container. View the gel on a UV transilluminator.
2. Make sure you wear UV-blocking goggles, gloves, and a lab coat or long sleeved shirt to protect yourself from the intense ultraviolet light emitted by the transilluminator. This will prevent a severe "sunburn" that can result from even short exposures to the UV light from the transilluminator.
3. A photograph of the gel will be taken using the photodocumentation system.
Laboratory Session 3
Analysis of Gel or Photograph
1. Examine the stained gel containing your sample and those from other individuals. Orient the gel with the sample wells at the bottom. First, ascertain whether you can see a diffuse (fuzzy) band of "primer dimer," that appears at the same position in each lane toward the top of the gel. Primer dimer is not amplified lambda DNA but is an artifact of the PCR reaction that results from primers amplifying themselves. Excluding primer dimer, interpret the bands in each lane of the gel:
2. Allele sizes can be estimated in agarose gels simply by comparing their positions to the ladder of size markers included in one lane of the gel. However, a closer determination of allele sizes can be obtained by graphing the function that determines the migration of linear DNA fragments through an agarose gel:
D =1/log MW
where D equals the distance migrated and MW equals the molecular weight of the fragment. For simplicity's sake, biologists substitute base-pair length for molecular weight in this calculation. .
3. Working from bottom (sample wells) to top of the gel assign the known base-pair sizes of the DNA standard to the bands appearing in the maker lane. The lambda/Hind III standard contains a 564-bp fragment that will run the fastest, followed by bands of 2027 bp and 2322 bp. Measure the distance (in mm) each marker fragment migrated from the sample well.
4. Set up semi-log graph paper with distance migrated as the X (arithmetic) axis and base-pair length as the Y (logarithmic) axis. Then, plot distance migrated versus base-pair length for each marker fragment. Connect data points with a line.
5. Measure and record distances migrated by the amplified lambda DNA fragment. To determine its base-pair size of an allele, first locate the distance it migrated on the X axis. Then, use a ruler to draw a vertical line from this point to its intersection with the marker data line. Now, extend a horizontal line from this point to the Y axis. The number on the Y axis is the calculated base-pair size of the allele.
6. How does the calculated size of the amplified lambda DNA fragment compare to its known size of 1106 bp?
ESTIMATION OF SIZE OF RESTRICTION FRAGMENTS BY REGRESSION ANALYSIS – GRAPHING IN MICROSOFT EXCEL
1. Open Microsoft Excel by clicking on the Excel icon.
2. Enter your data from your DNA size marker in Columns A (distance migrated) and Column B (fragment size).
3. After entering your data, click the Charting Icon on the toolbar, or select Chart in the Insert menu. The Chart wizard will open.
4. In the chart window select XY (Scatter) chart type. Continue by clicking Next.
5. Select the columns that you want to plot. Continue by clicking Next.
6. Select the Titles tab and fill in the information for the x-axis title as Distance (mm). Fill in the y-axis as Base Pairs (bp). Select the Gridlines tab and place checks for major and minor gridlines for both x- and y-axes. Select the Legend tab and remove the check by Show Legend. Continue by clicking Finish.
7. The graph should have been inserted into the spreadsheet. Place your cursor on the y-axis and double click. Select Scale tab and under the Value (Y) Axis Scale enter the size of the smallest fragment as Minimum and the size of the largest fragment as Maximum. Place a check next to Logarithmic scale. Hit OK.
8. Place your cursor of the x-axis and double click. Select Scale tab and under the Value (X) Axis Scale enter the distance the smallest fragment migrated as Minimum and the distance the largest fragment migrated as Maximum. Hit OK.
9. Next, add a trendline. Under Chart in the toolbar, select Add Trendline. In the trendline window, select exponential. Under Options, select R2 value and show equation. Click OK. Your graph should now show a linear line, regression equation, and R2 value. The regression equation shows the equation for the line generated by the data
10. Use the regression equation to estimate the size of your amplified PCR fragment.
SUGGESTED READINGS
Budowle, B., R. Chakraborty, A. M. Giuti, A. J. Eisenberg, and R. C. Allen. 1991. Analysis of the VNTR locus D1S80 by the PCR followed by high-resolution PAGE. Amer. J. Hum. Genet. 48:137-144.
Campbell, A. M., J. H. Williamson, D. Padula, and S. Sundby. Use PCR and a single hair to produce a “DNA fingerprint.” Amer. Biol. Teacher 59:172-178.
Mullis, K. 1990. The unusual origin of the polymerase chain reaction. Sci. Amer. 262:56-61.
Nakamura, Y., M. Carlson, K. Krapcho, and R. White. 1988. Isolation and mapping of a polymorphic DNA sequence (pMCT118) on chromosome 1p (D1S80). Nucleic Acids Res. 16:9364
Sajantila, A., B. Budowle, M. Strom, V. Johnson, M. Lukka, L. Peltonen, and C. Ehnholm. 1992. PCR amplification of alleles at the D1S80 locus: Comparison of a Finnish and a North American Caucasian population sample, and forensic casework evaluation. Amer. J. Hum. Genet. 50:816-825.
EXERCISE 13
CONTROL OF PROKARYOTE GENE EXPRESSION: LAC OPERON
EQUIPMENT
Laboratory Session 1
Water bath, 37oC
Laboratory Session 2
Spectronic 20 spectrophotometer
SUPPLIES AND MATERIALS
Laboratory Session 1
Lac+ strain of Escherichia coli (15 ml/group)
5 x 10-3 M Isopropyl-ß-D-thiogalactoside (IPTG)
0.5% Orthonitrophenyl-ß-B-galactoside (ONPG)
M Na2CO3
Toluene (1 dropper bottle/group)
5 ml pipettes
1 ml pipettes
Test tube racks (1/group)
Test tubes (10/group)
Laboratory Session 2
Spectrophotometer tubes (10/group)
Kimwipes
50 ml beaker (1/group)
Distilled water
ONPG SOLUTION (Store in Refrigerator)
Make up 100 ml phosphate buffer (pH 7.0) as follows:
60 ml Na2HPO4 . 2H2O (11.876 g/liter)
40 ml KH2PO4 (9.078 g/liter)
To every 100 ml of this solution add:
0.5 g ONPG
IPTG SOLUTION (Store in Refrigerator)
IPTG 0.12 g
Add 100 ml of distilled water
1.0 M SODIUM CARBONATE SOLUTION
Na2CO3 106.0 g
Water to 1.0 L
INTRODUCTION
In prokaryotes, regulation of transcription allows the cell to use its resources with great economy, which is important because there may be large fluctuations in the quantity of nutrients provided by the environment. The organization of the genome into operons means that manufacture of the products of adjacent genes--products that have sequential and related functions--can be turned on or off coordinately. For example, three enzymes are required for lactose utilization by Escherichia coli, and these three enzymes are coded for by three contiguous genes: lac z, lac y, and lac a (Fig. 1). The lac z gene specifies the amino acid sequence of the ß-galactosidase enzyme, which converts lactose to glucose and galactose. This conversion is essential if lactose is to serve as the primary energy source in glycolysis. The second gene, lac y, specifies the primary structure of ß-galactoside permease, which facilitates the entry of lactose into the bacterial cell. The third gene, lac a, codes for a transacetylase enzyme whose physiological role is unknown. These three structural genes are under the control of a regulatory gene designated lac i. The lac i gene regulates the transcription of the three structural genes by producing a repressor molecule which binds to the operator region of the operon and effectively represses transcription. However, when lactose is present in the medium, the repressor is prevented from binding to the operator and RNA polymerase binds to the promoter region of the operon and transcribes the structural genes. The RNA is then translated to produce the enzymes necessary for lactose metabolism (Fig. 2).
[pic]
Figure 1. The metabolism of lactose by ß-galactosidase to yield galactose and glucose.
Figure 2. Lactose operon of Escherichia coli.
In this experiment we will induce ß-galactosidase production in E. coli. We can easily assay for the presence of this enzyme if we use a substrate that undergoes a color change during the enzyme-catalyzed reaction. If the substrate and any necessary co-factors for the reaction are present in excess amounts, then the rate of the reaction (or color change) will be determined by the amount of enzyme present in the reaction mixture.
The enzyme ß-galactosidase catalyzes the hydrolysis of a wide variety of β-galactosides. The disaccharide lactose is a substrate and inducer for this enzyme. Isopropyl-β-D-thiogalactoside (IPTG) is a structural analogue of lactose and a more potent inducer (Fig. 3). Unlike lactose, IPTG is not hydrolyzed by the induced enzyme. To assay for the enzyme, another analogue of lactose, orthonitrophenyl-β-B-galactoside (ONPG), will be used. When this colorless compound is hydrolyzed, the intensely yellow nitrophenolate ion is produced (Fig. 4).
[pic]
Figure 3. Structure of lactose and Isopropyl-β-D-thiogalactoside (IPTG), which is a structural analogue of lactose.
[pic]
β-Galactosidase
Orthonitrophenyl-ß-D-galactoside __________________> Galactose + Orthronitrophenolate ion (Yellow at pH 8.0)
Figure 4. Hydrolysis of orthonitrophenyl-β-D-galactoside by ß-galactosidase.
OBJECTIVE
The experiment demonstrates the phenotypic behavior of a Lac+ strain of E. coli following growth in medium with isopropyl-β-D-thioglactoside (IPTG).
Laboratory Session 1
PROCEDURE
1. You will be given a 15 ml culture of E. coli. Pipet 5 ml of the culture to each of two tubes labeled 1 and 2. Add 0.6 ml of 10-3 M IPTG and 0.6 ml of distilled water to tube 1 and 1.2 ml of water to tube 2. Be sure to label each tube appropriately.
2. Mix the contents of the two tubes and as quickly as possible remove 1.0 ml from each of the two tubes. Put each sample in separate tube and label appropriately. Add a drop of toluene and shake vigorously. This is your 0 minutes sample.
3. Place the two tubes labeled 1 and 2 in the water bath at 37oC. After 20, 40, and 60 minutes of incubation repeat step 2 for tube 1. Make sure to add a drop of toluene to each sample.
4. After 60 minutes of incubation, repeat step 2 for the water treated culture.
5. When all samples have been taken, agitate each sample tube vigorously again and then add 0.2 ml of the ONPG solution to each sample. Shake again.
6. Place the sample tubes in a 37oC water bath for 20 minutes.
7. Add 2.8 ml of 1.0 M Na2CO3 to each sample tube to stop the reaction and intensify the color. The exact time interval between starting (addition of ONPG) and stopping (addition of Na2CO3) is recorded. The tubes should be stoppered and placed in a refrigerator until the next session.
Laboratory Session 2
PROCEDURE
Use of Spectrophotometer
The induction of β−galactosidase production will be measured using the spectrophotometer. Review the instructions for use of the Spectronic 20 prior to continuing the experimental protocol.
1. The parts of the Spectronic 20 spectrophotometer should be located with the help of Figure 1.
[pic]
Figure 1. Spectronic 20 spectrophotometer.
2. Power. Rotate the light source switch clockwise and allow the spectrophotometer to warm-up for 5 minutes.
3. Wavelength selection. Adjust wavelength control to 420 nm for this experiment.
4. Zero setting. Adjust the light source control until the meter reads 0% transmittance. NOTE: No test tube should be in the instrument and the cover of the sample holder must be closed.
Standardizing light control
1. Prepare a blank that contains 2.8 ml of 1.0 M Na2CO3, 0.2 ml of ONPG, and 1.0 ml of distilled water. Zero the spectrophotometer using the blank. Wipe fingerprints from the tube with a Kimwipe.
2. Insert the tube into the sample holder, aligning the mark on the spectrophotometer tube with the line on the sample holder. Close the cover.
3. Adjust the light intensity control until the meter reads 100% transmittance.
Sample measurement
1. Fill a spectrophotometer tube about 1/2 full of the sample to be measured. Do this for each sample.
2. Remove the tube of the blank solution and replace it with the tube containing the sample. Align the tube with the line on the sample holder. Close the cover.
3. Measure the optical density (O.D.) of each sample of supernatant at 420 nm. If the absorbance of a sample exceeds the scale on the spectrophotometer, dilute the sample 1:10 for your OD measurement and then multiply the reading times 10.
4. Plot the O.D. 420 against the minutes of incubation for all cultures.
SUGGESTED READINGS
Maniatis, T. and M. Ptashne. 1976. A DNA operator-repressor system. Sci. Amer. 234:64
Ptashne, M. and W. Gilbert. 1970. Genetic repressors. Sci. Amer. 222:36-44.
APPENDIX A
INSTRUCTIONS FOR JOURNAL STYLE LABORATORY REPORTS
INSTRUCTIONS FOR JOURNAL STYLE LABORATORY REPORTS
Laboratory reports will be prepared in the format of a scientific paper and should follow the style of journals such as Genetics or Journal of Heredity. Examine papers in recent issues of these journals to familiarize yourself with details of organization, section headings, methods of data presentation and reference citations. A scientific paper has six sections: Title, Abstract, Introduction, Materials and Methods, Results, Discussion, and Literature Cited. The various sections of the scientific paper and their contents are described below. It is often easiest to start with your Materials and Methods section or Results section. Once these sections are written, proceed to the discussion followed by the Introduction, Title, Literature Cited, and finally Abstract.
Format for Scientific Paper
Title
The title should concisely describe the main topic of the paper and usually contains fewer than ten words. Your title should be self-explanatory when standing alone. Do not use abbreviations in the title, and all words should be spelled out for clarity. The organism used in the study can be identified in the title using its scientific name. Remember that both components (genus and specific epithet) are underlined or italicized (Drosophila melanogaster or Drosophila melanogaster). Capitalize only the genus name. Avoid superfluous phrases such as "Study of", "Investigations on".
Abstract
The abstract is a one-paragraph summary of your research. It should include what was done, general procedures, main findings, and conclusions. The latter should be limited to those statements that can be directly inferred from the results. Write the abstract in the past tense. The abstract should be the last section of the paper written.
Introduction
The introduction includes background information on methods used, on previous research in the same area that is pertinent to your research, on historical aspects of the experiments, and on anything that needs to be explained so that a person who was unfamiliar with the topic could understand the purpose and importance of your research. The purpose of your research should be included in the Introduction, usually in a sentence or short paragraph at the end of the section. Any nonoriginal information contained in this section (or any other section) must be cited.
In the body of your paper (Introduction, Materials and Methods, Discussion), you should cite references by the author’s last name and year. If there are three or more authors, you should cite the first author’s last name followed by “et al.” (the Latin et al. means “and others”).
Examples:
1. One author – Lane, 1995
2. Two authors – Lane and Jones, 1995
3. Three or more authors – Lane et al., 1995
Materials and Methods
Describe in detail the material, equipment, and methods in paragraph format. Do not present these items in the form of a list. This section should be explicit enough to allow someone to repeat your experiments. A chronological pattern should be used, and this section should be written in the past tense. When procedures from a lab book or another paper or report are followed exclusively, simply cite the work.
Results
The data obtained from your research is presented in this section, in a logical and unbiased manner. Data should not actually be interpreted in the Results section--just present what happened, but not why (save the whys for the Discussion). A well-written Results section will include a paragraph or two at the beginning to briefly explain how the data were obtained. Your actual results should be presented in the form of an appropriately labeled table or figure. Numbers or other hard data should be shown in a table (See Table 1 as an example). Graphs or diagrams are shown as a figure (see Figure 1 as an example). The text should not repeat what is evident from the figures and tables but should point out the salient features and should connect the results with one another. Write in the past tense.
Figures and tables should be self-explanatory; that is the reader should be able to understand them without referring to the text of your paper. All columns and rows in tables and axes in figures should be labeled.
Guidelines for Tables (See example below):
1. All tables should be referred to in the text of your paper (e.g. Table 1).
2. Each table should be typed on a separate sheet of paper.
3. Each table should have a title above the table that describes its content.
4. All tables are numbered consecutively (e.g. Table 1, Table 2, etc).
5. Double-space the contents of a table.
Table 1. Restriction fragments sizes
determined from the digestion of lambda
DNA with Hind III
|Fragment |Size (kb) |
| | |
|1 |23.13 |
|2 |9.42 |
|3 |6.56 |
|4 |4.36 |
|5 |2.32 |
|6 |2.03 |
Guidelines for a figure (See example below)
1. All figures should be referred to in the text of your paper (e.g. Figure 1 or Fig. 1)
2. Each figure is typed on a separate sheet of paper.
3. All figures are numbered consecutively.
4. A caption that describes the content of the figure should be placed underneath the figure.
5. Axes on graphs should be labeled properly to included units of measure when applicable.
6. In a graph, the dependent variable is plotted on Y-axis and the independent variable on the X-axis.
[pic]
Figure 1. Standard curve generated from the digestion of lambda DNA with the restriction enzyme Hind III.
Discussion
In the Discussion section, the data presented in the Results section is interpreted and discussed. Your discussion should emphasize interpretation of data and relate the data to existing theory and knowledge. Compare the results of your study to those of other studies and offer suggestions for differences. The most important thing to remember about writing a discussion is not to reiterate the results. A well-written Discussion includes the following:
1. What were the major points illustrated by the data? Present principles, relationships, and generalizations shown by results. Discuss do not repeat results.
2. Do your results agree with previously published work? The previously published work must be referenced in the Literature Cited section. Point out exceptions and unexplained results and define unsettled points. Show how your results and interpretations agree (or disagree) with previously published work.
3. Is the data contradictory in itself?
4. Discuss theoretical implications or practical applications of the work.
5. Clearly state your conclusions.
6. Summarize your evidence for each conclusion.
7. The discussion should end with a short summary or conclusion regarding the significance of the work.
Literature Cited
Any information you obtain from a source other than yourself must be referenced. Footnoting or endnoting is not an acceptable means of citing literature for scientific writing. Each piece of literature cited should be listed and alphabetized by first author. The following is a widely used means of referencing other work:
Author, author's initials, date of publication (year), title of paper or book, name of journal with its volume and inclusive pages, or name and city of publisher with number of pages used in book.
Examples
Journal Article:
Marmur, J. 1961. A procedure for the isolation of deoxyribonucleic acid from micro-organisms. J. Mol. Biol. 3:208-218.
Book:
Sambrook, J., E.F. Fritsch, and T. Maniatis. 1989. Molecular Cloning: A Laboratory Manual, 2nd ed., Cold Spring Harbor Laboratory Press, NY, pp. 100-108.
General Guidelines for Writing Scientific Paper
1. Underline or italicize scientific names (Drosophila melanogaster or Drosophila melanogaster).
2. Write out the scientific name of an organism initially and abbreviate afterward (Drosophila melanogaster then D. melanogaster).
3. Use metric system of measurements. Abbreviate units without a period (e.g. ml, µl, g, etc).
4. Numbers are written as numerals when they are greater than ten or when associated with measurements. Never start a sentence with a numeral.
5. Start each paragraph with a topic sentence.
6. Make sure each sentence has a subject and a verb and that they are in agreement.
7. Avoid using the first person. Use third person and past tense when you are writing about an experiment that has already been conducted. Active voice is usually more readable.
8. Make sure pronouns refer to their antecedents.
Guidelines for Scientific Paper
1. Laboratory reports must be typed. Use 12 point font (Times, Helvetica, or Geneva) to write your paper.
2. Your paper must include a minimum of three references. These references can include textbooks, reference manuals, and journal articles.
3. At least one of your references must be a journal article.
4. Include a photocopy of each journal article used in your paper. Photocopy the page(s) of any other reference used and include with your paper.
APPENDIX B
INSTRUCTIONS FOR MAINTAINING A LABORATORY NOTEBOOK
INSTRUCTIONS FOR MAINTAINING A LABORATORY NOTEBOOK
Each student is required to keep a laboratory notebook. The notebook should be a complete record of the experimental work performed in the laboratory. Your entries should be thorough and well organized and should permit another person to understand the experiment and repeat the work in precisely the same manner in which the original experiment was done. Thus, accuracy, completeness, and clarity in your records are of primary importance. Your laboratory notebook will be an invaluable record and should contain all the information needed to write your scientific papers for the course.
General Guidelines:
1. Obtain a bound Mead composition notebook from the University Bookstore. Spiral bound or loose-leaf notebooks are not acceptable.
2. Put your name, the course number, and the laboratory section on the outside front cover of the notebook.
3. All entries in your notebook are to be made in non-erasable ink with printing preferred for clarity. Never use "white-out" in your notebook. If you make a mistake, never obliterate or write over the mistake. Draw a single line through the mistake and make the correction above, below, or beside the error.
4. Before using the notebook, subsequent right hand pages should be consecutively numbered in ink. Never tear pages from your notebook. If a page is to be ignored or voided, simply draw a diagonal line through the page.
5. Reserve the first two pages in your notebook for a Table of Contents. The Table of Contents should be filled out as the laboratory assignments are completed. The Table of Contents should refer to the page number of the first page of each experiment.
6. Do not write on the back of pages. Put a corner-to-corner diagonal line through them to void the blank spaces.
7. Start every new experiment on a fresh page. Any unused portions of a page at the end of an experiment should have a diagonal live drawn through it.
8. If your experiment lasts over several laboratory periods, the notebook should be dated each time you work on the project.
9. The maintenance of a satisfactory laboratory notebook is extremely important. To help you learn to maintain a laboratory notebook, notebooks will be graded periodically during the semester. All experiments must be recorded in your laboratory notebook, and a maximum of ten (10) points will be awarded for each laboratory experiment. The time of the first grading session will be announced in advance; however, all subsequent grading sessions will not be announced in advance.
Format for Laboratory Notebook
There are several acceptable formats for a laboratory notebook. For this course, we will use the format outlined below:
1. Date at the top of each page
2. Title of Experiment
3. Name(s) of laboratory partner(s)
4. Purpose
5. Procedure
6. Observations and Data
7. Calculations
For each experiment, organize your notebook into sections beginning with the headings given above. This will facilitate recording and locating information regarding the experiment.
Pre-Laboratory Events
1. Read the laboratory exercise so that you are thoroughly familiar with the experiment.
2. Prior to coming to the laboratory, you should have made the following entries in your laboratory notebook regarding the experiment:
3. Title of experiment - This is the title of the experiment given in the laboratory manual
4. Purpose - In your own words, state the purpose of the experiment in a concise and succinct manner. The purpose should indicate what you are investigating and what is your experimental approach. Do not simply copy the purpose given in the laboratory manual.
5. Procedure - It is not necessary to copy the detailed procedure given in the laboratory manual; however, you should provide a reference to the source(s) of the procedure. For example: The procedure followed the experimental design described in Laboratory Exercises in Genetics by M. Beck, Fall 1999, pp. 50-55.
Laboratory Events
1. Record the current date and names of any lab partners you may have for this experiment. The date should be recorded at the top of every page.
2. Procedure - Note any changes to the procedure given in the laboratory manual.
3. Observations and Data - Your experimental data and observations are to be directly recorded in the notebook in ink at the time they are obtained. Never record information on scraps of paper. Record units with all numbers that have units, e.g. µl, g, etc.
4. Use tables to display data whenever possible. Tables should follow the format suggested in the laboratory manual. The table should have a descriptive title, and each column of the table should be labeled showing what quantity is displayed and in what units.
5. Data can also be displayed using graphs. Each axis of the graph should have numerical values, units, and a label as to what it represents. The graph should have a descriptive title. When plotting data the independent variable, or quantity being varied, should be plotted on the horizontal axis (abscissa). The dependent variable, or quantity being measured, should be plotted on the vertical axis (ordinate).
6. Observations are a critical and often overlooked part of the laboratory notebook. Include anything noteworthy that you observe such as cloudy culture medium, precipitate, temperature changes, or unexpected problems with the equipment or procedures.
7. Affix any computer spreadsheet, graph, photograph, or similar information obtained during the experiment directly into the notebook with transparent tape. An adequate description should accompany each item. Partners may photocopy original data for inclusion in the lab notebook.
Post-Laboratory Events
1. Show all calculations associated with the experiment, no matter how simple they may seem.
2. When performing calculations, show any equations or formulas used. Be certain that all symbols are adequately defined and that all answers have the appropriate units.
APPENDIX C
REGRESSION ANALYSIS
REGRESSION ANALYSIS
Scientists use an experiment to search for cause and effect relationships in nature. In other words, they design an experiment so that changes to one item cause something else to vary in a predictable way. These changing quantities are called variables. The independent variable is the one that is changed by the scientist. As the scientist changes the independent variable, he or she observes what happens. The dependent variable changes in response to the change the scientist makes to the independent variable. The new value of the dependent variable is caused by and depends on the value of the independent variable.
Regression is a technique use to predict the value of a dependent variable using one or more independent variables. There are two types of regression analysis namely simple and multiple regressions. Simple regression involves two variables, the dependent variable and one independent variable. Multiple regression involves many variables, one dependent variable and many independent variables. Mathematically, the simple regression equation is as shown below:
y = α + βx
In simple linear regression, a single dependent variable, Y, is considered to be a function of an independent X variable, and the relationship between the variables is defined by a straight line. (Note: many biological relationships are known to be non-linear and other models apply.) When a best-fit regression line is calculated, its binomial equation defines how the variation in the X variable explains the variation in the Y variable.
When an investigator has collected two series of observations and wishes to see whether there is a relationship between them, he or she should first construct a scatter diagram. The vertical scale represents one set of measurements and the horizontal scale the other. If one set of observations consists of experimental results and the other consists of a time scale or observed classification of some kind, it is usual to put the experimental results on the vertical axis (Y axis). These represent what is called the "dependent variable". The "independent variable", such as time or height or some other observed classification, is measured along the horizontal axis (X axis), or baseline.
The regression equation representing how much y changes with any given change of x can be used to construct a regression line on a scatter diagram, and in the simplest case this is assumed to be a straight line. The direction in which the line slopes depends on whether the correlation is positive or negative. When the two sets of observations increase or decrease together (positive) the line slopes upwards from left to right; when one set decreases as the other increases the line slopes downwards from left to right. As the line must be straight, it will probably pass through few, if any, of the dots. Given that the association is well described by a straight line we have to define two features of the line if we are to place it correctly on the diagram. The first of these is its distance above the baseline; the second is its slope. They are expressed in the following regression equation where β is the slope and α is the intercept (the point where the line crosses the y axis):
[pic]
[pic]
If ( < 0 the line slopes downward X and Y are inversely related.
If ( > 0 the line slopes upward X and Y are positively related
The amount of variation in the dependent variable that is accounted for by variation in the predictor variables is measured by the value of the coefficient of determination, often called R2 adjusted. The coefficient of determination, R2, is a measure of how well the variation of one variable explains the variation of the other, and corresponds to the percentage of the variation explained by a best-fit regression line which is calculated for the data. The closer this is to 1 the better, because if R2 adjusted is 1 then the regression model is accounting for all the variation in the outcome variable. As the value of R2 increases, one can place more confidence in the predictive value of the regression line.
Analysis and Sizing of DNA Fragments
In a charged field, DNA fragments migrate through a matrix (gel) in roughly a size dependent manner, which is described by a log-linear equation. We can use this relationship to calculate the sizes of unknown DNA fragments separated by agarose gel electrophoresis. In order to determine the size of DNA molecules, we need to compile data of the molecular size as a function of the migration distance and make a graph using a semi-log scale for the y-axis of a standard DNA (i.e. one digested with a restriction enzyme and whose fragment sizes are known). Such a standard curve can be compiled using Microsoft Excel and regression analysis. The regression equation can then be used to calculate the size of DNA fragments whose migration distances in the gel are known.
Open an Excel worksheet and input the migration distance measurements (independent variable) in Column A and molecular sizes (dependent variable) from the ladder lane (standard DNA) on the agarose gel in Column B (see example below). Once you have your data entered, you can create a graph of the distance vs the fragment length. Finally, a regression equation (regression model has to be selected. This is referred to as adding a “trimline” in Excel.
|[pic] | | | | |
Excel offers a choice of five different equations:
|Linear regression |y = a + b x |
|Logarithmic regression |y = a + b ln(x) |
|Exponential regression |y = a bx |
|Power regression |y = a xb |
|Polynomial regression |y = a + bx + cx2 |
Choose the “Exponential” graph, which will display your results in semi-log form with which you are familiar. You now have both a graphical representation of the relationship between the size and migration distance of your DNA fragments, and the mathematical description of that relationship (regression equation). Since you can measure X (migration distance) for any fragment on the gel, you can calculate accurately the value of Y using the regression equation.
Example: Suppose you have a fragment on the gel that migrated 50 mm. You can insert this value into the regression equation and solve for the size of the DNA fragment.
y = 80659e-0.0753x (regression equation from graph)
y = 80659e(-0.0753 * 50)
y = 1869 base pairs
-----------------------
Figure 1. One letter symbol for each amino acid
Black strain Tan strain
[pic]
R2 Adjusted
Regression Equation
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